Cyanogenic glycosides: synthesis, physiology, and phenotypic plasticity.

PP65CH06-Gleadow

ARI

ANNUAL REVIEWS

8 April 2014

21:52

Further

Annu. Rev. Plant Biol. 2014.65:155-185. Downloaded from www.annualreviews.org by Universita degli Studi di Roma La Sapienza on 05/17/14. For personal use only.

Click here for quick links to Annual Reviews content online, including: • Other articles in this volume • Top cited articles • Top downloaded articles • Our comprehensive search

Cyanogenic Glycosides: Synthesis, Physiology, and Phenotypic Plasticity Roslyn M. Gleadow1 and Birger Lindberg Møller2,3 1 School of Biological Sciences, Monash University, 3800 Victoria, Australia; email: [email protected] 2 Plant Biochemistry Laboratory, Department of Plant and Environmental Sciences; Center for Synthetic Biology “bioSYNergy”; and VILLUM Research Center for Plant Plasticity, University of Copenhagen, DK-1871 Copenhagen, Denmark 3

Carlsberg Laboratory, DK-1799 Copenhagen, Denmark

Annu. Rev. Plant Biol. 2014. 65:155–85

Keywords

First published online as a Review in Advance on February 24, 2014

cyanogenesis, hydrogen cyanide, cytochrome P450, plant defense, pathogen interactions, herbivore interactions

The Annual Review of Plant Biology is online at plant.annualreviews.org This article’s doi: 10.1146/annurev-arplant-050213-040027 c 2014 by Annual Reviews. Copyright All rights reserved

Abstract Cyanogenic glycosides (CNglcs) are bioactive plant products derived from amino acids. Structurally, these specialized plant compounds are characterized as α-hydroxynitriles (cyanohydrins) that are stabilized by glucosylation. In recent years, improved tools within analytical chemistry have greatly increased the number of known CNglcs by enabling the discovery of less abundant CNglcs formed by additional hydroxylation, glycosylation, and acylation reactions. Cyanogenesis—the release of toxic hydrogen cyanide from endogenous CNglcs—is an effective defense against generalist herbivores but less effective against fungal pathogens. In the course of evolution, CNglcs have acquired additional roles to improve plant plasticity, i.e., establishment, robustness, and viability in response to environmental challenges. CNglc concentration is usually higher in young plants, when nitrogen is in ready supply, or when growth is constrained by nonoptimal growth conditions. Efforts are under way to engineer CNglcs into some crops as a pest control measure, whereas in other crops efforts are directed toward their removal to improve food safety. Given that many food crops are cyanogenic, it is important to understand the molecular mechanisms regulating cyanogenesis so that the impact of future environmental challenges can be anticipated.

155

PP65CH06-Gleadow

ARI

8 April 2014

21:52

Contents


INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . STRUCTURE AND BIOSYNTHESIS OF CYANOGENIC GLYCOSIDES . . . . . . Structural Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Turnover . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . CYANOGENESIS AND THE CYANIDE BOMB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . HERITABILITY, POLYMORPHISM, AND QUANTITATIVE VARIATION . . . . Presence/Absence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Quantitative Inheritance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Where and When: Age Versus Developmental Stage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . HERBIVORES, PATHOGENS, AND WOUNDING RESPONSES. . . . . . . . . . . . . . . Herbivory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pathogen Resistance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wounding and Induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ENVIRONMENTAL PLASTICITY AND OPTIMAL GROWTH . . . . . . . . . . . . . . . . Effects of Nitrogen Supply . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Atmospheric CO2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nonoptimal or Stressful Growth Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . RESOURCE ALLOCATION: PURPOSES, COSTS, AND REGULATION . . . . . . . Energy Costs and Trade-Offs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nitrogen as the Currency for Calculating Costs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

156 158 158 159 161 162 163 163 165 165 166 166 167 168 169 169 170 171 172 172 174

INTRODUCTION

CNglc: cyanogenic glycoside Cyanogenesis: the release of HCN from endogenous natural products containing a stabilized α-hydroxynitrile functional group

156

Humans have a complicated relationship with hydrogen cyanide (HCN): It is toxic, and yet plants that generate HCN are an important part of the human diet. All plants produce tiny amounts of HCN as an additional product in the biosynthesis of ethylene, but some plant species can release large amounts from endogenously stored cyanogenic glycosides (CNglcs). CNglcs are bioactive natural products (also called specialized plant products or secondary metabolites) derived from amino acids with oximes and cyanohydrins (α-hydroxynitriles) as key intermediates. Glucosylation of the labile cyanohydrin results in the formation of a CNglc (133). CNglcs are stable compounds, but when the β-glycosidic linkage is hydrolyzed through the action of a β-glycosidase, the labile cyanohydrin that is formed dissociates to release HCN in a process known as cyanogenesis (137) (Figure 1). Acute or chronic exposure to HCN can lead to intoxication, mild to severe illness, and in extreme cases even death in humans (120) and animals (51) because HCN inhibits the activity of metalloenzymes, principally cytochrome c oxidase, the final enzyme in the respiratory electron transport chain (115). Hydrolysis of CNglcs results in the concomitant release of carbonyl compounds that may further augment the toxic and repellant effects of HCN (156). The oxime intermediates in the biosynthesis of CNglcs may also be toxic, especially toward fungi (99, 132). CNglcs and the specific catabolic enzymes required to catalyze their hydrolysis are stored in separate compartments within cells or in different tissues, preventing accidental autotoxicity (137). The two components are brought into contact upon tissue disruption, as caused by chewing herbivores or when cell integrity is destroyed by physical processes, such as by freezing or maceration Gleadow

·

Møller

PP65CH06-Gleadow

ARI

8 April 2014

21:52

Amino acid

Oxime P450aa

P450ox

α-Hydroxynitrile (cyanohydrin)

HCN

R2

UDP-glucose

Glucose

O Sugar

UDPglucosyltransferase

α-Hydroxynitrile

C R1

N β-Glucosidase

Cyanogenic monoglucoside

α-Hydroxynitrile lyase (or spontaneous) Aldehyde/ketone


Figure 1 Biosynthesis and bioactivation of cyanogenic glycosides (CNglcs). CNglcs are synthesized from specific amino acids in a series of reactions catalyzed by two multifunctional, membrane-bound cytochrome P450s (P450aa and P450ox) and a soluble UDP-glucosyltransferase, with an oxime and an α-hydroxynitrile (cyanohydrin) as key intermediates. Cyanogenesis occurs when the β-glucosidic linkage is hydrolyzed by a specific β-glycosidase to form an unstable α-hydroxynitrile (cyanohydrin) that dissociates into hydrogen cyanide (HCN) and a ketone either spontaneously at high pH or catalyzed by an α-hydroxynitrilase. Abbreviations: aa, amino acid; ox, oxime.

of the plant material as part of food processing (70). The cyanide potential (HCNp) is defined as the total amount of HCN that could be released by complete conversion of all cyanogenic compounds in a given tissue to HCN (39); the cyanogenic capacity (HCNc) is the catabolic rate of HCN release from CNglcs (15). CNglcs are effective deterrents to generalist herbivores (17, 70, 190, 204), and this is most likely the main evolutionary driver in their occurrence across the plant kingdom (3, 9, 124). In some plant species, CNglcs have been implicated in the control of germination (52, 162) and bud burst via cyanohydrin and HCN formation (20). CNglcs may also serve as transport forms of carbon and nitrogen (179), and endogenous turnover processes may release the nitrogen from CNglcs in the form of ammonia (90, 140, 157). More recently, it has been proposed that CNglcs may also function in modulating oxidative stress (98, 133, 140). CNglcs are unusually widespread in the plant kingdom. They are found in the oldest of terrestrial plants (the ferns) and in gymnosperms and angiosperms (205). More than 3,000 plant species from all major vascular plant taxa, belonging to over 130 families of flowering plants, are cyanogenic; these represent 11% of the total number of plant species tested, with a clear overrepresentation of cultivated plants. This proportion is, therefore, somewhat higher than the 5% found in screens of natural population systems (65, 128). Throughout history, humans appear to have inadvertently selected cyanogenic plants for cultivation, perhaps because of their increased resistance to herbivores or because the need to process the plant material before consumption makes them less attractive to trespassers (93, 124). The focus of this review is on the formation, turnover, and degradation of CNglcs and how their contribution to phenotypic plasticity may optimize adaptation to environmental challenges. The synthesis and regulation of bioactive natural products are influenced by changes in the CO2 content of the atmosphere (62, 64) and by changes in the climate (31, 194). It is important to be aware of and understand the consequences of such changes so that appropriate measures can be taken to either reduce risks or capitalize on benefits because, for example, increased levels of CNglcs may alter the host specificity of insects or pests. This is highly relevant for CNglcs, considering the option to bioengineer CNglc production into naturally noncyanogenic plant species in order to improve their resistance to herbivores and pests as an alternative to the use of pesticides and fungicides (56, 190). Attempts are also being made to use classical mutagenesis-based breeding techniques, or RNA-interference or antisense technology, to reduce the toxicity of edible tissues of crop plants that accumulate large amounts of these compounds in order to make food crops safer for consumption (25, 95). In nature, some plant species, such as white clover (Trifolium repens), are www.annualreviews.org • Cyanogenic Glycosides

Cyanide potential (HCNp): the maximum amount of HCN that could be released from a given tissue; it provides an alternative measure of the total CNglc concentration Cyanogenic capacity (HCNc): the catabolic rate of HCN release from CNglcs; it is dependent on the total CNglc content and the activity of the catabolic enzymes β-glycosidase and α-hydroxynitrilase

157

PP65CH06-Gleadow

ARI

8 April 2014

21:52

clearly polymorphic with respect to their ability to produce and degrade CNglcs. CNglcs are one of many types of modulators of plant plasticity, and under most environmental conditions they are not necessarily essential components. Given the known roles of CNglcs and their integration into plant primary metabolism, such manipulations may result in unexpected effects over the course of plant ontogeny (33). New understandings of the diverse roles of CNglcs shed light on the physiological mechanisms underlying their contribution to improved phenotypic plasticity in addition to their classical role as a constitutive defense system. At the end of the review, we list key knowledge gaps and directions for future research on CNglcs.

STRUCTURE AND BIOSYNTHESIS OF CYANOGENIC GLYCOSIDES Annu. Rev. Plant Biol. 2014.65:155-185. Downloaded from www.annualreviews.org by Universita degli Studi di Roma La Sapienza on 05/17/14. For personal use only.

Structural Diversity All of the CNglcs so far identified are derived from aromatic and aliphatic amino acids (tyrosine, phenylalanine, valine, isoleucine, and leucine) and a few nonproteinogenic amino acids: the (2S,1 R) and (2S,1 S) epimers of 2-(2 -cyclopentenyl)-glycine as well as (2S,1 S,2 S)-2-(2 -hydroxy3 -cyclopentenyl)-glycine (36, 39, 133, 177). The most common CNglcs are the monoglucosides dhurrin [(S)-4-hydroxymandelonitrile-β-D-glucopyranoside], prunasin [(R)-mandelonitrile-β-Dglucopyranoside], linamarin (2-hydroxyisobutyronitrile-β-D-glucopyranoside), and lotaustralin [(R)-2-hydroxy-2-methyl-butyronitrile-β-D-glucopyranoside] (Figure 2), which are derived from tyrosine, phenylalanine, valine, and isoleucine, respectively. Cyanohydrins derived from tyrosine, phenylalanine, and 2-(2 -cyclopentenyl)-glycine (and notably not valine) contain a chiral center, giving rise to pairs of epimeric CNglcs like dhurrin and taxiphyllin, prunasin and sambunigrin, and deidaclin and tetraphyllin A, respectively (149). The core structures of CNglcs may be further modified by single or multiple hydroxylations. These decorations are particularly common among the isoleucine- and leucine-derived CNglcs, such as those found in barley (143) and the genetic model plant Lotus japonicus (53, 187). Hydroxylation may also result in cleavage of the aromatic ring system, as observed in the biosynthesis of the tyrosine-derived triglochinin (142). It has not been resolved whether the hydroxylated CNglcs with a cyclopentanoid core structure are derived from hydroxylated parent amino acids or whether hydroxylation proceeds as a late step in their synthesis (36). The number of CNglcs found in nature is further expanded by the structural diversity of the sugar moiety. In the CNglcs currently known, the cyanohydrin is always stabilized by a βglucosidic linkage to D-glucose. In cyanogenic diglycosides, the second sugar moiety may also be a D-glucose residue, with the most common examples being amygdalin (prunasin-6 -glucoside) and linustatin (linamarin-6 -glucoside), in both of which the second glucose residue is attached by a β-1,6 linkage. However, the second glucose residue may also be bound in β-1.2, β-1.3, and β-1.4 linkages, as in eucalyptosin A, C, and B, respectively (139). Further structural diversity arises by incorporation of, e.g., arabinose, xylose, and apiose residues as additional sugar residues, as observed in vicianin, xeranthin, lucumin, and oxyanthin (4, 129, 163, 175). Yet another level of diversity arises from additional combinations of acylation and glycosylation reactions, as observed in oxyanthin 5 -benzoate (163). The presence of these derivatives is typically restricted to specific stages in plant ontogeny or to specific tissues (129, 131). Isolation and structural characterization are thus challenging and dependent on advanced liquid chromatography–mass spectrometry and NMR instrumentation, so only within the past few decades has the widespread occurrence of these complex derivatives of the classical CNglcs been fully realized. Rather than being considered a rather small class of specialized molecules, CNglcs are now thought of as a rapidly expanding and diverse group of compounds. 158

Gleadow

·

Møller

PP65CH06-Gleadow

ARI

8 April 2014

21:52

OH H

OH O

HO HO

CN

O

HO HO

O

OH

a

CN

H

OH

O

HO HO

O

O

O

OH

OH

b

Prunasin

CN

H

O OH HO HO

c

Sambunigrin

Amygdalin

OH CN

OH O

HO HO

O

HO HO

CN O

OH HO HO

O

OH


d

O

HO HO

g

HO HO

O

CN

CN

O

HO HO

O

Lotaustralin H

O HO HO

O

O O

OH

O

O

OH

f

Linustatin

OH H

OH

O

OH

e

Linamarin

CN

OH

O

O

h

Epiheterodendrin

OH

Tetraphyllin B

i

O

OH

OH OH

OH

CN

Oxyanthin 5''-benzoate

OH H

OH HO HO

O

CN O

j

k

Proteacin

O

HO HO

OH

O

H

OH

OH

HO

HO HO HO

O

O

HO HO

OH OH

O

OH

NC

OH

HO

O

O

OH

l

Triglochinin

CN

OH

Dhurrin

OH O

O

O HO

O

OH

O O

O OH HO HO

H O

CN OH

O

OH

OH OH

m

O

Xeranthin

Figure 2 The structural diversity of cyanogenic glycosides (CNglcs). Prunasin (a) and sambunigrin (b) are epimers derived from phenylalanine. Amygdalin (c) is a prunasin-derived diglucoside. Linamarin (d ) and linustatin (e) are mono- and diglucosides (respectively) derived from valine. Lotaustralin ( f ) is derived from isoleucine and typically co-occurs with linamarin. Epiheterodendrin ( g) is derived from leucine. Tetraphyllin B (h) is derived from the nonproteinaceous amino acid 2-cyclopentenyl glycine following an additional hydroxylation. Oxyanthin 5 -benzoate (i ) is an example of a more complex CNglc derived from phenylalanine and containing glucose and apiose residues. Proteacin ( j ), triglochinin (k), and dhurrin (l ) are all derived from tyrosine. Xeranthin (m) is one of the most structurally complex CNglcs identified so far, being an acylated tetraglycoside derived from phenylalanine following an additional 3-hydroxylation. The sugar residues involved are glucose, xylose, and apiose, the latter of which is acylated by a cinnamic acid residue.

Biosynthesis The biosynthetic pathway for CNglcs was first elucidated in Sorghum bicolor (122, 134, 182). In sorghum, the CNglc dhurrin is synthesized from L-tyrosine in a series of steps catalyzed by two multifunctional cytochrome P450 enzymes (CYP79A1 and CYP71E1) (8, 99, 100, 109, 183) and a family 1 UDP-glucosyltransferase (UGT85B1) (94) in combination with the P450 redox partner NADPH-dependent cytochrome P450 oxidoreductase (POR) (79) (Figure 3). The www.annualreviews.org • Cyanogenic Glycosides

159

160 H2N

Gleadow

·

Møller

NH3 +

N NH2

HO O

O

H2 S

OH

O OH

H2N NH2

N

OH

or

HO NH2

N

L-Aspartic acid

O

O OH

Dhurrin

N

Glucose

Glucose

Alternate turnover pathway

O

Glutathione conjugate

Glutathione

Glutathione S-transferase?

GlutathioneS-lyase

+ NH3

Glucose

HO

Catabolism

UGT85B1

UDP-glucose

Dhurrinase

p-Hydroxyphenylacetonitrile

HO

OH

L-Asparagine

2H2O

O

O

HO

N

OH

p-Hydroxymandelonitrile

HO

p-Hydroxymandelonitrile

NADP+

CYP71E1

Nitrilase heteromer NIT4

Nitrilase

OH

NADPH + O2

α-Hydroxynitrile lyase

(E)-p-Hydroxyphenylacetaldehydeoxime

p-Hydroxyphenylacetic acid

HO

N

p-Hydroxybenzaldehyde

HCN +

CYP79A1

β-Cyanoalanine

β-Cyanoalanine synthase

HO

NADP+

8 April 2014

Cysteine

Detoxification

L-Tyrosine

OH

O

NADPH + O2

ARI

Scheme illustrating the synthesis and turnover of dhurrin in Sorghum bicolor. The steps in the putative alternative turnover pathway resulting in the conversion of dhurrin into 4-hydroxyphenylacetonitrile have not been defined (90, 157), but it is known that SbNIT4A/B2, a nitrilase heterodimer, catalyzes the conversion of 4-hydroxyphenylacetonitrile to 4-hydroxyphenylacetic acid and ammonia (90). p-Hydroxyphenylacetic acid glucoside has been shown to accumulate in etiolated sorghum seedlings (78, 90).

Figure 3

HO

Biosynthesis


PP65CH06-Gleadow 21:52


PP65CH06-Gleadow

ARI

8 April 2014

21:52

first step is the conversion of tyrosine to (E)-p-hydroxyphenylacetaldoxime (183). This step, catalyzed by CYP79A1, is rate limiting in the biosynthesis of dhurrin in young sorghum seedlings (32). The (E)-p-hydroxyphenylacetaldoxime is then converted to p-hydroxymandelonitrile in a CYP71E1-catalyzed reaction (8). Glucosylation of p-hydroxymandelonitrile catalyzed by the soluble UGT85B1 enzyme generates dhurrin (94). p-Hydroxymandelonitrile is labile at neutral and alkaline conditions, and if it is not rapidly glucosylated, it dissociates into HCN and phydroxybenzaldehyde. The enzymes catalyzing dhurrin formation in sorghum (CYP79A1, CYP71E1, UGT85B1, and POR) are thought to form a metabolon to promote rapid channeling of the toxic and labile intermediates into dhurrin formation and to prevent undesired metabolic crosstalk (91, 97, 113, 144). CYP79A1, CYP71E1, and POR are anchored to the endoplasmic reticulum via a transmembrane domain positioned near their N termini (8, 109), whereas UGT85B1 is a soluble enzyme (94). Metabolon assembly may be mediated by direct protein–protein interactions or by specific lipids (91, 190). It remains to be understood how the glycosides are transported from their initial production site in the cytosol not only to their storage site, typically in the vacuole (169), but also to the specific tissues such as the epidermal cell layer and epicuticular wax of the barley leaf (88, 143). The types of intermediates identified in the CNglc pathway in sorghum have also been found in other species, such as flax (Linum usitatissimum) (44), white clover (T. repens) (87), cassava (Manihot esculenta) (6), L. japonicus (53), almond (Prunus amygdalus) (167), and Triglochin maritima (142). Thus, the key intermediates involved in biosynthesis of CNglcs appear to be the same across the plant kingdom. In addition to sorghum, the genes encoding the entire pathway for CNglc synthesis have now been identified in cassava (6, 96, 105) and L. japonicus (188). Genes encoding CYP79s and UGTs involved in CNglc synthesis have also been identified in white clover (150) and almond (57), respectively. The aliphatic CNglcs linamarin and lotaustralin normally co-occur in plants, but the ratios between the two CNglcs differ between species. The cassava genome contains two paralogs of each of the three biosynthetic genes involved in linamarin and lotaustralin formation. Preliminary analysis shows identical regulation and coexpression of the different genes and their paralogs (96, 105). In L. japonicus, the paralogous genes CYP79D3 and CYP79D4 show differential expression, with CYP79D3 preferentially expressed in areal parts of the plant and CYP79D4 expressed exclusively in the roots (53). The CYP79s are considered the signature enzyme of the CNglc pathway, and all tested current members of this P450 family catalyze conversion of amino acids to the corresponding oximes. Both CYP79 and the second P450 in the pathway, catalyzing the conversion of oxime to cyanohydrin, belong to the CYP71 clan of P450s (7). The latter P450 and the UGT that catalyzes the final step in CNglc synthesis have a much broader substrate specificity compared with that of CYP79 (80, 94, 100).

Turnover In plants containing large amounts of CNglcs, the nitrogen and sugar content of the CNglcs may represent significant percentages of the total organic matter present (140). For example, in newly formed shoots of Eucalyptus cladocalyx, as much as 25% of leaf nitrogen can be tied up in prunasin (64); in tips of young etiolated sorghum seedlings, dhurrin constitutes 30% of the dry matter (78); and in the epidermal cell layer of barley leaves, glucose bound in CNglcs constitutes 90% of the total soluble carbohydrate content (160). Under such circumstances, but also in general, the ability of plants to reuse the carbon and nitrogen deposited in CNglcs (e.g., to balance resource demands in primary metabolism) offsets some of the energy and resource costs associated with their synthesis and storage (140). www.annualreviews.org • Cyanogenic Glycosides

161

ARI

8 April 2014

21:52

The classical catabolic pathway proceeds after tissue disruption and involves hydrolysis by a β-glycosidase and dissociation of the cyanohydrin formed by the action of an α-hydroxynitrilase (39, 86) (Figures 1 and 3). This pathway is typically referred to as the bioactivation pathway or cyanogenesis. Part of the HCN may be recaptured and reincorporated into primary metabolism in a reaction catalyzed by β-cyanoalanine synthase that involves stoichiometric consumption of cysteine (26, 158). Subsequent action of nitrilase 4 (NIT4) family enzymes results in asparagine and aspartate formation (90). More recently, an alternative pathway independent of prior tissue damage was discovered and studied in sorghum (90). p-Hydroxyphenylacetonitrile is an intermediate in this turnover pathway and is converted into p-hydroxyphenylacetic acid by the action of the nitrilase heteromer NIT4A/NIT4B2s with a concomitant release of ammonia (90) (Figure 3). The turnover rates achieved by this mechanism may be quite high, as demonstrated in young sorghum plants, where it may reach 0.8% of the total CNglc present per hour (1). Rapid turnover of dhurrin is also observed upon extended growth of sorghum plants (32).


PP65CH06-Gleadow

CYANOGENESIS AND THE CYANIDE BOMB The process resulting in the release of HCN from CNglcs by the action of a β-glycosidase is referred to as cyanogenesis, and sometimes also as the cyanide bomb (137). Hydrolysis of glucosinolates by the action of myrosinases represents a similar binary system, in this case resulting in the release of thiocyanates and related compounds, generally referred to as the mustard oil bomb (205). In both these binary systems, compartmentalization is essential to prevent autotoxicity and to ensure that the “bomb” is detonated as a targeted response, e.g., to herbivore attack (137). The most common type of compartmentation is at the level of the organelle. CNglcs, at least in the leaves, are typically confined to the vacuole (e.g., 40, 98, 169). Within the stems and petioles of cassava, linamarin is confined to vesicle-like structures within the latex (48). The location of the β-glycosidases involved in hydrolysis of CNglcs is more variable. Depending on the species, they may be present in the apoplastic space, bound to the cell wall, in the cytoplasm, in small vesicles, or in the chloroplast (48, 58, 101, 111, 165, 191). Less is known about the location of the α-hydroxynitrilases, but the evidence available suggests that they are cytoplasmic in both sorghum (111) and Hevea brasiliensis (84). Separation of CNglcs and their degradative enzymes may also occur at the tissue level. In sorghum, for example, dhurrin is found almost exclusively in the epidermal cell layer of the leaf blade, whereas dhurrinase and α-hydroxynitrilase are located predominantly in the mesophyll cells (111, 169, 191). A similar epidermal localization of CNglcs has also been observed in lima beans (Phaseolus lunatus) (58). In barley (Hordeum vulgare), the CNglcs and β- and γ-hydroxynitrile glucosides are likewise localized in the epidermal cells of the leaf blade, whereas β-glucosidase activity related to their hydrolysis is restricted to the endosperm of the germinating seed (143). Within fruits, some plants concentrate the CNglcs in the seeds [e.g., H. brasiliensis (179), Prunus serotina (186), and Canthium schimperianum (174)], whereas in others, the cyanogenic component is the fruit itself [e.g., Passiflora edulis (34)]. Some species have only trace amounts of CNglcs in the dry seeds [e.g., sorghum (120) and clover (87, 195)], whereas others have completely acyanogenic seeds [e.g., E. cladocalyx (69)]. Prunasin and amygdalin are located in the parenchyma of the cotyledons in the seeds of rosaceous stone fruits, whereas the β-glucosidase and α-hydroxynitrilase are located in the procambium (159, 165, 186). The β-glycosidases responsible for hydrolysis of CNglcs have been isolated and their genes identified in several species. Sorghum contains two β-glucosidases, the dhurrinases Dhr1 and Dhr2, which hydrolyze dhurrin with high specificity and organ-specific expression (35). Dhr1 accumulates in the mesocotyl and the root tip, whereas Dhr2 accumulates in the leaves (35, 191), 162

Gleadow

·

Møller


PP65CH06-Gleadow

ARI

8 April 2014

21:52

although the functional significance of this difference is unknown. White clover contains a single linamarase encoded by the Li locus (152). L. japonicus contains the two paralogous β-glucosidases BGD2 and BGD4. BGD2 is able to hydrolyze all the nitrile glucosides present in L. japonicus, including linamarin and lotaustralin. In contrast, BGD4 specifically hydrolyzes the isoleucinederived β- and γ-hydroxynitrile glucosides designated as rhodiocyanoside D and A, respectively (136, 188). The β-glucosidases discussed above have cyanogenic monoglucosides as substrates. Bitter almonds contain large amounts of the cyanogenic diglucoside amygdalin and smaller amounts of the monoglucoside prunasin (167). Amygdalin is hydrolyzed by amygdalin hydrolase, resulting in the formation of the monoglucoside prunasin, which may then be cleaved by a prunasin hydrolase. Amygdalin hydrolase and prunasin hydrolase constitute small enzyme families (110). No evidence for formation of the diglucoside gentiobiose was observed upon the action of amygdalin hydrolase (167). In contrast, β-glycosidase-catalyzed cleavage of the cyanogenic diglycoside vicianin results in formation of the diglycoside vicianose, composed of arabinose and glucose (4). Thus, depending on the plant species and CNglc, β-glycosidase-catalyzed hydrolysis of CNglcs harboring more than a single sugar moiety may proceed sequentially as well as by direct initial release of the aglycone. Cyanogenic diglycosides are frequently referred to as storage or transport forms. Formation of the diglycosides may prevent their hydrolysis by β-glycosidases, as documented in H. brasiliensis, where the linamarase hydrolyzing the monoglucoside linamarin was shown to not cleave the co-occurring diglucoside linustatin (76).

HERITABILITY, POLYMORPHISM, AND QUANTITATIVE VARIATION Presence/Absence Cyanogenesis is a stable, heritable trait. In some species, all individuals are cyanogenic, whereas others are highly polymorphic, with phenotypically acyanogenic individuals (e.g., 42, 54, 67, 87, 130, 170, 202). Relatively few species are monomorphic (e.g., 95, 121, 125, 128). Cyanogenesis in white clover is a classic example of Mendelian inheritance with two independently segregating loci—one indicative of the ability to produce CNglcs (Ac) and the other indicative of the presence of the specific β-glucosidase able to degrade the CNglcs (Li ) (43, 81, 87, 151, 152). Plants must be homozygous recessive at one of the loci to be acyanogenic. Both phenotypes persist in the population as a consequence of selection for both cyanogenic and acyanogenic phenotypes under different environmental and herbivore pressures, giving rise to a balanced polymorphism (43, 87, 104). Plants with the Acaclili genotype are not naturally cyanogenic, although they are still able to synthesize CNglcs. Plant material containing CNglcs but without enzymes able to degrade them may pose a toxicity risk because the CNglcs may be broken down to release HCN in some animals, such as in cattle rumens (51). For example, Turnera ulmifolia may contain the specific β-glucosidase even when the capacity to synthesize CNglcs is absent (170), whereas Acacia species that are known to synthesize CNglcs always lack the requisite β-glucosidase for natural cyanogenesis (41). Although the genetics of cyanogenesis has been studied in clover for more than 50 years, the actual basis of the mutation was only recently established: The nonfunctional ac allele is the result of a deletion in CYP79D15, the gene encoding the first P450 in the pathway (151). Lack of linamarase is linked to partial or complete loss of the Li gene from the white clover genome (152). There is no evidence of linkage between Ac and Li in clover at the phenotypic (87) or molecular level (151, 152), despite the obvious advantages of inheriting both together. A recently described ethyl methanesulfonate–generated mutation in sorghum, tcd1, that renders CYP79A1 nonfunctional is also recessive and follows a Mendelian pattern of inheritance (25), also without any indication of www.annualreviews.org • Cyanogenic Glycosides

163

Manihot esculenta


Sorghum bicolor

Lotus japonicus

PP65CH06-Gleadow

ARI

8 April 2014

21:52

CYP79D4

UGT85K3

Rho

CYP736A2

CYP79D3

CYP736p

CYP736p

Chromosome 3, contig CM0241 (region 140–560 kb)

CYP736p

20 kb

CYP71E1

UGT85B1

CYP79A1

CYP71

Chromosome 1 (region 1,040–1,170 kb)

10 kb

CYP79

CYP71E7 CYP79D2

Scaffold 08265 (region 850–950 kb)

UGT85K4

CYP71E

UGT85K5

CYP71E and CYP736 UGT85

10 kb

Figure 4 Schematic representation of the clustering of cyanogenic glucoside biosynthetic genes in the genomes of Lotus japonicus (chromosome 3), Sorghum bicolor (chromosome 1), and Manihot esculenta (chromosome number not yet established because the genome is still in draft form). Functional genes are shown with arrows indicating their orientation. Confirmed genes in cyanogenic glucoside biosynthesis are labeled above each bar, with CYP79 genes in pink, CYP71E and CYP736 genes in green, and UGT85 genes in blue. Adapted from Reference 187.

linkage. The last step in cyanogenesis is the dissociation of the cyanohydrin with concomitant release of HCN. This process is catalyzed by an α-hydroxynitrilase but occurs spontaneously at alkaline pH. Little is known about the α-hydroxynitrilases catalyzing the dissociation process (24). In L. japonicus, sorghum, and cassava, the three genes coding for the specific enzymes that catalyze the formation of CNglcs are clustered on a single chromosome (187, 189) (Figure 4). Clustering of nonhomologous genes encoding enzymes for synthesis of plant defense compounds is a recently discovered phenomenon. It was first reported in maize (Zea mays) for the synthesis of the benzoxazinoid defense compound DIMBOA (2,4-dihydroxy-1,4-benzoxazin-3-one) (61). Some terpenoid pathways are also organized in gene clusters. This applies to synthesis of avenacin in oat (Avena strigosa) (138, 161), synthesis of momilactone and multifunctional phytocassanesoryzalides/oryzadiones in rice (Oryza sativa) (196), and synthesis of steroidal glycoalkaloids in potato (Solanum tuberosum) and tomato (Solanum lycopersicum) (89). The genes encoding synthesis of the opiate alkaloid noscapine in the opium poppy (Papaver somniferum) are also clustered (201). In CNglc biosynthesis, the individual enzymes catalyzing a specific step are encoded by genes that are related between species but not necessarily by orthologous genes. Likewise, the organization of the three known gene clusters encoding CNglc synthesis in the three species investigated is very different (187). Together, these point to independent evolution of CNglc biosynthesis in several higher-plant lineages by repeated recruitment and neofunctionalization of members from similar multigene families (187, 189). Some insects are able to synthesize cyanogenic glucosides de novo (202, 203), as observed in the Burnet moth (Zygaena filipendulae). The insect genes CYP405A2, CYP332A3, and UGT33A1, which encode the entire biosynthetic pathway, have been recruited 164

Gleadow

·

Møller

PP65CH06-Gleadow

ARI

8 April 2014

21:52

from the same two multigene families as in plants. The low sequence identity between the plant and insect genes clearly demonstrates that the insect pathway has evolved by convergent evolution and not following horizontal gene transfer or by divergent evolution, as might have been assumed considering that the P450s involved are multifunctional and catalyze highly unusual reactions (72, 92). Thus, plants and insect have independently found a way to package a cyanide time bomb to fend off herbivores, predators, and pests. Interestingly, the genes coding for the cyanogenic βglucosidases are not clustered with the biosynthetic genes (187, 189), consistent with the observed independence between β-glucosidase activity and HCNp in nonmodel species (67, 172).


Quantitative Inheritance CNglc concentration (HCNp) shows high quantitative variability even within genetically identical plants of single clones (31, 193). Broad-sense heritability, calculated by comparing the cyanogenic potential of half sibs, has been found to be approximately 0.8 in several species (e.g., 19, 72, 173). Given that the inheritance of the trait is well known, this is likely to be due to other factors, such as differences in physiology or in uptake and allocation of resources. The activity of cyanogenic β-glycosidases varies quantitatively as well and appears to be a determining factor in the rate of HCN release from CNglcs (HCNc) (11). A strong positive correlation between HCNp and β-glucosidase activity has been reported in Eucalyptus polyanthemos (73) but was not detected in natural populations of E. cladocalyx (68).

Where and When: Age Versus Developmental Stage One of the difficulties in comparing the cyanogenic status of individual plants is that the HCNp varies ontogenetically, phenologically, and chronologically. HCNp is generally highest in seedlings and decreases with plant age (45, 69, 114, 193, 199). For example, in E. cladocalyx, in the series Sejunctae, seedlings have a high HCNp (74) (Figure 5). Notable exceptions to the pattern described above are the cyanogenic Eucalyptus species from the series Maidenaria. They are essentially acyanogenic as seedlings ( 600 ppm) (200). At some stages of plant ontogeny, like the seedling stage, the HCNp is likely transcriptionally regulated (32), whereas at other stages regulation may be controlled at multiple levels. A link between nitrogen supply and CNglc deployment has also been observed in legumes, where the rate of colonization by nitrogenfixing rhizobia has been associated with higher concentrations of linamarin and lotaustralin and decreased herbivory in both clover (106) and lima beans (14). Not all plants respond to nitrogen in this way. In a study by Busk & Møller (32) (see above), dhurrin concentration did not increase in very young seedlings grown at high levels of potassium nitrate. Moreover, transcript levels Downregulation of RuBisCO

R2 O

Shade

Sugar R1

C N

CNglc

Young tissue (growth)

Extreme temperatures

Improved water use efficiency?

CO2

Nitrogen

Promotion of new growth


Wounding induced a surge in HCN release in lima beans and Hevea by increasing β-glucosidase and α-hydroxynitrilase activity rather than increasing HCNp (11, 98). Herbivores may elicit a different response to mechanical damage mediated through chemical signals or enzyme activities in their saliva, as has been found in Nicotiana attenuata (46). Many plants increase methyl jasmonate and salicylic acid levels in response to herbivory and pathogen attack (49, 168). In sorghum, microarray studies have revealed extensive transcriptional changes in response to wounding or exposure to methyl jasmonate and salicylic acid (47, 164); these changes include induction of dhurrinase and α-hydroxynitrilase transcripts and temporary induction of CYP71E1, one of the genes governing dhurrin biosynthesis (208), similar to the enzyme activity change in lima beans and Hevea described above. Rather than being a constitutively expressed defense system, CNglc biosynthesis and cyanogenesis obviously respond to environmental and developmental changes.

Retardation of growth

Drought Acceleration of maturation

Figure 8 Scheme illustrating environmental plasticity in cyanogenic glycosides (CNglcs). CNglcs are typically higher in plants where growth is limited by resources other than nitrogen. Conditions associated with an increase in CNglcs are shown as arrows, those associated with a decrease in CNglcs are shown as lines with circles on the end, and those where the effect is uncertain are indicated with a dashed line. Interplays between the different parameters, as they are thought to affect CNglcs, are also indicated (Table 1). www.annualreviews.org • Cyanogenic Glycosides

169

PP65CH06-Gleadow

ARI

8 April 2014

21:52

Table 1 Environmental effects of cyanogenic glycoside (CNglc) concentration on plasticity Functional group


Species

Nitrogen

Phosphorus

Drought

Elevated CO2

Sorghum bicolor, Sorghum sudanense

C4 grass

Incr. (32, 121, 154, 200)

Decr. (154, 200)

Incr. (147, 200)

NS (66)

Trifolium repens

Legume

Incr. (106)

Decr. (195)

Incr. (81, 195)

NS (58, 62)

Shade

Incr. (195)

Lotus corniculatus

Legume

NS (27)

Phaseolus lunatus

Legume

Incr. (12)

Pteridium esculentum

Fern

Manihot esculenta

Euphorb

Incr. (63)

Ryparosa kurrangii

Rainforest understory tree

NS (199)

NS (199)

Prunus turneriana

Rainforest understory tree

NS (127)

NS (127)

Eucalyptus cladocalyx

Woodland tree

Incr. (69, 184)

Eucalyptus polyanthemos

Woodland tree

NS (72, 73)

Eucalyptus yarraensis

Woodland tree

NS (75)

High temperature

Incr. (185, 195)

NS (22) Incr. (12) Incr. (42) Incr. (125, 194)

Incr. (71)

NS (63)

NS (64)

Decr. (28)

Decr. (30)

Conclusions are based on data from controlled-environment or agricultural experiments. Frequencies of cyanogenic phenotypes in populations reflect evolutionary responses and are not included here. CNglc concentration increased (Incr.), decreased (Decr.), or was not significantly different (NS) in plants grown experimentally at higher levels of nutrients (nitrogen or phosphorus), with low soil moisture (drought), in elevated CO2 , in shade, or at a high temperature. Figure 8 shows possible interdependencies. Species are listed by their functional groups.

for CYP79A1 and CYP71E1 were unchanged, presumably because production was already at its maximum. Controlled-environment studies of E. polyanthemos, Prunus turneriana, and R. kurrangii seedlings also found no correlation between nitrogen supply and foliar HCNp (72, 127, 199). The exact reasons for this are not clear but may be related to particular life history traits. Seedlings of P. turneriana and R. kurrangii seedlings, which remain suppressed in the rain-forest understory for many years, have high background levels of defense compounds and, like young sorghum seedlings, may be at maximal expression. By contrast, HCNp is barely detectable in young E. polyanthemos seedlings, where activation of the pathway may be dependent on plant ontogeny (139).

Atmospheric CO2 Improved nitrogen use efficiency of plants growing at higher concentrations of atmospheric CO2 presents the possibility of a reallocation of nitrogen to CNglcs (33). Controlled experiments using clover (59, 62), E. cladocalyx (64), and cassava (tubers) (63) did not detect any direct effect of CO2 on the molar concentration of accumulated CNglcs. Experiments that measured the different nitrogen pools, however, detected a shift in the allocation of nitrogen: Leaves had a significantly lower concentration of protein, resulting in a higher ratio of CNglcs to protein (62, 64) (Figure 9). From a nutritional point of view, the relative proportion of protein and CNglc content is important because sulfur amino acids are required for the detoxification of released HCN (Figure 6). 170

Gleadow

·

Møller

PP65CH06-Gleadow

ARI

8 April 2014

a

21:52

b Prunasin RuBisCO Other nitrogen sinks (nucleic acids, etc.) Other protein


Ambient CO2 (380 ppm)

Twice ambient CO2 (780 ppm)

Figure 9 Allocation of nitrogen to prunasin [measured as cyanide potential (HCNp)], RuBisCO, total other protein, and other nitrogen sinks (such as nucleic acids and thylakoid membranes) in the leaves of Eucalyptus cladocalyx seedlings grown at either (a) ambient or (b) approximately twice the ambient concentrations of atmospheric CO2 . The area of each pie is proportional to the total nitrogen concentration of the leaves in each treatment (64).

Nonoptimal or Stressful Growth Conditions CNglc concentrations are generally higher when growth is limited by environmental factors such as light, temperature, or drought (Figure 8, Table 1). Three explanations are often presented to account for this: (a) CNglcs are concentrated in a smaller amount of plant tissue (178), (b) the plants are phenologically younger owing to delayed growth (126), or (c) there is active upregulation at the transcriptional level (32, 47, 164, 208). Active upregulation could be either an adaptation to protect existing plant tissue from herbivores or a mechanism for moderating oxidative stress (133, 178). Transcriptomic analysis of sorghum seedlings experiencing osmotic stress found more than 1,000 genes with altered expression, including many involved in defense-related pathways (29, 47). Abscisic acid, an important signal molecule in drought responses and regulation of stomatal aperture, also covaries with HCNp (5, 18). Evidence for a direct effect of stress on CNglc synthesis comes from the study of cell cultures. Osmotically stressed suspension cultures of Eschscholtzia californica showed higher production rates of dhurrin and triglochinin (85), which suggests that the synthesis of these compounds was induced by the osmotic stress. The magnitude of the increase in HCNp in response to low soil moisture depends on the severity and duration of the stress, the ontogenic stage, and the availability of other resources (71, 147, 194). In E. cladocalyx, the HCNp of extant leaves increased by 70% after a prolonged drought when the nitrogen supply was high but only by 30% when the nitrogen supply was limited (70). Similar responses in cyanogenic crop plants may have important ramifications for human health (see sidebar, Cyanide and Cassava). In cassava, drought-stressed tubers may become more toxic because of a direct increase in concentration (194) and relocation of linamarin from leaves to tubers (103). This increased HCNp in drought-stressed cassava is not permanent and decreases after plants are rewatered (194). A similar situation exists in forage sorghum, where a severe or extended drought period may result in toxic levels of dhurrin and necessitate long-term silage to enable use of the sorghum plants as fodder (25, 147). CNglcs may also provide additional plasticity to plant development by controlling seed germination in response to rainfall events. Eremophila maculata grows in arid regions, and the fruit walls of its seeds contain germination inhibitors in the form of water-soluble glycosides, including www.annualreviews.org • Cyanogenic Glycosides

171

PP65CH06-Gleadow

ARI

8 April 2014

21:52


CYANIDE AND CASSAVA A single dose of cyanide of ∼1–3 mg per kilogram of body weight is lethal for most vertebrates (148). Cassava has a high HCNp, and it is the only staple food that is potentially lethal if not adequately processed (124). The HCNp of cassava flour bought at markets in Mozambique in a year suffering from drought was greater than 200 ppm (50), well above the World Health Organization’s recommended maximum level of 10 ppm. Under such special circumstances, where water required for proper processing is limited and the population is restricted to a monotonous cassava diet high in cyanohydrins or cyanide, ingestion of the toxic constituents may cause acute cyanide poisoning or development of konzo, a neurological disease causing permanent paralysis of the lower limbs (124). The concentration of the CNglcs linamarin and lotaustralin increases in cassava tubers when the plant suffers from severe drought (124). Elite cassava lines that yield well in different soil types and when subjected to different environmental challenges concomitant with having a low HCNp are in high demand (95).

the CNglc eucalyptosin A. Heavy rainfall causes the CNglc to leach out of the fruit wall and thereby trigger germination under optimal growth conditions (162). In rosaceous plants (e.g., almonds), cold acclimation and bud break and flower development may be controlled by cyanohydrin or HCN release resulting from endogenous hydrolysis of the CNglcs present (20). The same mechanisms may explain why spraying with cyanamide (which causes HCN liberation) can induce and synchronize bud break and flowering in some rosaceous plants. Only a few studies have considered the effect of temperature on HCNp. In T. repens, HCNp is typically higher in plants growing at nonoptimal temperatures, e.g., below 15◦ C or above 25◦ C (185, 195). Foliar HCNp has been found experimentally to be higher, lower, or unaffected in plants grown in the shade (e.g., in clover, E. cladocalyx, and P. turneriana, respectively) (30, 81, 127). The different responses appear to reflect the environments to which the species are adapted. In E. cladocalyx, which is adapted to open woodlands, nitrogen was reallocated from prunasin production to the photosynthetic apparatus in shade-grown plants, likely to maximize light interception and increase CO2 fixation (30). In contrast, P. turneriana seedlings, adapted to long periods under a rain-forest canopy, reallocated CNglcs away from growing leaf tips to the existing leaves, presumably to protect existing productive leaves from herbivory (127).

RESOURCE ALLOCATION: PURPOSES, COSTS, AND REGULATION Energy Costs and Trade-Offs Resource allocation trade-off theories assume that there must be a cost associated with synthesizing and maintaining defensive compounds (38, 83). For CNglcs, this has been hard to demonstrate, leading some to speculate that the costs are too small to measure (107). The average energy cost of dhurrin synthesis has been calculated as 2.1 g glucose (g dhurrin)−1 (60). This is low in terms of the overall energy budget, although additional costs would have to be assigned to maintenance, turnover, and transport as well as the synthesis of the β-glucosidase and α-hydroxynitrilase required for rapid HCN release. The bottom line is that the cost of deploying a CNglc-based defense must be less than the benefit gained, e.g., through reduced grazing (81) or increased metabolic plasticity. If there are costs in CNglc deployment, then it should be possible to measure a sacrifice in terms of plant growth or reproductive output. Highly cyanogenic E. polyanthemos seedlings were found to grow more slowly than seedlings with low HCNp (72). However, in a similar study using E. cladocalyx seedlings in which ontogenetic effects were kept to a minimum, no trade-off 172

Gleadow

·

Møller


PP65CH06-Gleadow

ARI

8 April 2014

21:52

was detected between CNglc concentration and relative growth rate or net assimilation rate (184). In clover, a small negative trade-off between CNglc synthesis and growth has been observed (55), but reproduction appears to be more strongly impacted (104). Cyanogenic clover morphs with the Ac genotype produce fewer flowers when grown in the absence of herbivory, but the difference in reproductive output is out of proportion to the energy costs involved. The difference in energy between Acac (low-flowering) and acac (high-flowering) phenotypes in terms of linamarin is only approximately 5 kJ, whereas the difference in terms of flower production is 130 kJ (102). The energy costs of synthesizing and maintaining CNglcs may be offset by their evolutionarily acquired roles in other parts of the metabolism and the use of shared pathways and metabolic crosstalk (140) (Figure 10). Synthesis and turnover of secondary metabolites such as CNglcs may be a way of dissipating excess energy and reducing power, thereby mitigating stress (107, 133, 141, 178, 180) (Figure 10). Synthesis of CNglcs consumes NADPH, directly dissipating any excess reducing power produced during light capture. CNglcs may also be able to quench reactive oxygen species, resulting in their conversion into amides, which, following hydrolysis, would liberate the Assembled metabolon Disassembled metabolon Amines

Amides

Amino acid

Mitochondrial dysfunction Camalexin Oxime

Detoxification

Reactive oxygen species Pathogens

Phytoalexins Indole-3-acetic acid

Metabolism as a response to biotic challenges Cyanogenic glucosides

Herbivore defense

Hydrogen cyanide Aldehyde or ketone

Endogenous catabolism

Glucose

Nitrile

Carboxylic acid

Primary metabolism

Ammonia Hydrogen cyanide

β-Cyanoalanine

Aspartic acid Asparagine

Figure 10 Mechanisms for minimizing biosynthetic cost and proposed multiple roles for cyanogenic glycosides (CNglcs). Costs are minimized using existing pathways. CNglcs also have secondarily acquired roles (e.g., in transport, defense, and possible stress tolerance), further offsetting the costs. Adapted from Reference 140. www.annualreviews.org • Cyanogenic Glycosides

173

ARI

8 April 2014

21:52

nitrogen in the form of easily accessible ammonium (180). The proposed alternative turnover pathway (157) (Figure 3) also results in amide formation. CNglcs may also react chemically with H2 O2 , opening up the possibility that the pathway helps the plant to reduce damage caused by excess reactive oxygen species (90, 133, 180). CNglcs are only one of many defenses at a plant’s disposal. Defense strategies are likely to vary with different selective pressures (magnitude and type) and with developmental stage (2, 17). Attempts to understand the complete defense syndrome are an increasingly important area for research (2). Trade-offs between strategies are likely, e.g., between CNglcs and tannins (27, 74) or volatile organic compounds (13). The higher HCNp in younger plants and plant parts is consistent with the optimal allocation theory of plant defense (123), but as leaves expand, there may simply be a trade-off with leaf toughness and other forms of chemical defense (198). New techniques under development—such as in situ polymerase chain reaction (95, 166), Raman spectroscopy (192), Fourier transform infrared spectroscopy (82), and bioimaging using mass spectrometry (117)—will likely help pinpoint localization and answer these questions.


PP65CH06-Gleadow

Nitrogen as the Currency for Calculating Costs Given that nitrogen is the limiting plant nutrient in most terrestrial systems, it may be more meaningful to calculate costs in terms of nitrogen. An analysis of E. cladocalyx trees sampled across a discontinuous natural population indicated that, on average, for every nitrogen molecule effectively incorporated into the leaves as prunasin, additional nitrogen was added on top of this investment, possibly to increase the capacity to generate additional energy by photosynthesis (68, 184). The roles of CNglcs in nitrogen storage and in defense are not necessarily mutually exclusive. The reduced rate of photosynthesis in stressed plants makes it economically attractive for plants to store nitrogen in a reduced form, ready to be remobilized when conditions improve, rather than as photosynthetic proteins or nitrate (147). In H. brasiliensis, the carbon and nitrogen resources in CNglcs can, for example, be mobilized and used for growth and latex production (112). Some species appear to have become obligate cyanotypes that are dependent on CNglcs for normal development at specific stages of ontogenesis. Cassava plantlets engineered using RNAinterference technology to have a highly reduced CNglc content exhibit poor growth or lethality at the time point where the wild-type plant would start to produce CNglcs and have a higher requirement for nitrogen supply (95). In a similar fashion, acyanogenic tcd1 seedlings of sorghum, in which the CYP79A1 protein harbors a single amino acid change that inactivates the enzyme, exhibit a slower initial growth rate than their wild-type siblings (25). In several species, studies have documented the ability of cyanogenic plants to remobilize the nitrogen bound to CNglcs into ammonia and its subsequent use in primary metabolism in the synthesis in amino acids without the release of gaseous HCN (37, 118, 125, 176) (see Figure 3). The ability of plants to remobilize the sugar and nitrogen stored in CNglcs, combined with the ability to transport CNglcs to specific parts of the plants guided by the generation of differently glycosylated transport forms, illustrates how turnover of CNglcs may play an important role in balancing primary metabolism, especially during major changes in plant ontogeny, and thereby improve plant plasticity (e.g., 133, 159, 176, 178). SUMMARY POINTS 1. Cyanogenic glycosides (CNglcs) are glycosides of cyanohydrins that release toxic hydrogen cyanide (HCN) and ketones when hydrolyzed by β-glycosidases and αhydroxynitrilases in a process referred to as cyanogenesis.

174

Gleadow

·

Møller

PP65CH06-Gleadow

ARI

8 April 2014

21:52

2. The biosynthesis of CNglcs and the process of cyanogenesis are now well understood in several species, and the relevant genes have been identified. 3. CNglcs are a structurally much more diverse class of bioactive products than originally thought, encompassing additional and different sugars than glucose that are linked by multiple linkage types and further decorated by acylation reactions.


4. Cyanogenesis is an effective defense against generalist herbivores but is not particularly effective against fungal pathogens. Many fungi efficiently convert HCN into ammonia and carbon dioxide. Some insect specialists have evolved mechanisms to sequester or de novo synthesize CNglcs and use them as their own defense against predators and as a source of reduced carbon and nitrogen. 5. Cyanogenesis follows classical Mendelian inheritance patterns, but its quantitative expression at a particular point in space and time is highly plastic. 6. The genes encoding the enzymes involved in CNglc biosynthesis are clustered to promote stable inheritance of the defense pathway in dynamic ecosystems. 7. CNglc concentration is generally higher in young plants, where growth is constrained, or where nitrogen is in ample supply. 8. Factors affecting CNglc concentration can be explained in terms of a resource-based trade-off between plant growth and defense. The difficulty in calculating such costs may arise because the production costs are actually low and because CNglcs have secondarily acquired important roles in nitrogen transport and storage and offer improved tolerance to oxidative stress, offsetting the direct costs of production. 9. CNglcs are defense compounds that through evolution have gained additional functions in plant primary metabolism manifested in the course of plant ontogeny. Many plant species are polymorphic with respect to cyanogenesis, demonstrating that CNglcs are not essential plant constituents but rather constituents that under specific environmental conditions and ontogenetic stages improve phenotypic plasticity by increasing the ability of plants to adapt to environmental challenges. Given the high proportion of food crops that are cyanogenic, it is important to be able to predict the response to changing environments in order to maintain and improve food safety.

FUTURE ISSUES 1. Most of our knowledge of CNglcs has been derived from studying a limited number of plant species. Recurrent evolution of the biosynthetic pathway has recently been demonstrated. In a similar manner, the routes for endogenous turnover may be highly species dependent. Likewise, the functional roles that CNglcs may play in a particular plant species to improve plant plasticity toward a particular environmental challenge and in the course of plant ontogeny may vary substantially. This also applies to the ability of CNglcs to ward off herbivores and pests and thus improve plant defense. Even at the most basic level, we need to take into account whether knowledge of the roles of CNglcs in one plant species can be generalized or whether the observed effects are unique.

www.annualreviews.org • Cyanogenic Glycosides

175

PP65CH06-Gleadow

ARI

8 April 2014

21:52

2. Many basic questions remain about the interaction of CNglcs with primary metabolism and their ability to facilitate plant plasticity in a variety of settings, including in response to environmental challenges. New roles for CNglcs are being proposed and need further scrutiny. Knowledge of the molecular mechanisms—e.g., with respect to increased stress tolerance—may be used to improve other crop plants through classical or molecular breeding. Given that so many food crops are cyanogenic, understanding the effects of climate change on CNglc accumulation and turnover is important for human food safety. Likewise, climate change–induced alterations in CNglc content may severely affect the feeding behavior of herbivores and pests, changes that could be either beneficial or harmful (or neither) with respect to humans and the global environment. Annu. Rev. Plant Biol. 2014.65:155-185. Downloaded from www.annualreviews.org by Universita degli Studi di Roma La Sapienza on 05/17/14. For personal use only.

3. Experimental data in the literature are frequently confounded by developmental and size differences. There is a need for additional experimentation that focuses on ontogenetically relevant comparisons and takes into account the differences in abiotic and biotic challenges that plants are exposed to in the course of their life cycles (126). Experiments in which the biological variation among plants is minimal may provide helpful guidelines on how to design large-scale field experiments (126). 4. The molecular regulation of CNglc production and turnover is poorly understood. Comparisons of the transcriptomes of acyanogenic and cyanogenic morphs, coupled with the metabolomics of the different nitrogen pools, would allow investigation of the physiological changes that accompany phenotypic plasticity. 5. The consequences for plant growth and agroecology of bioengineering cyanogenesis into and out of crop plants—as an alternative to the use of pesticides and to reduce their toxicity, respectively—need to be carefully evaluated. Cassava is propagated by cuttings, so the negative effects of removing CNglcs at the seedling stage may not have any practical effects in cassava cultivation, but careful investigations to identify negative effects in the course of plant ontogeny need to be strengthened and encouraged. 6. Several studies have indicated that there may be a trade-off between cyanogenesis and other types of defense. The emergence of new model species with mutations in different parts of the biosynthetic and catabolic pathways as well as mutant collections in crop plants presents unique opportunities to discover how the potentially conflicting demands of responding to abiotic stress, herbivory, and attack by pathogens can be reconciled. 7. Use of crops for green manure as a substitute for chemical fertilizers and pesticides is an important approach for more sustainable agricultural practices. Green manure from cyanogenic crops like white clover and sorghum is rich in nitrogen and releases HCN that may be utilized for pest control. The risk of leaching HCN into the environment following application to sandy and loamy soils needs to be investigated to avoid unintended threats to the quality of groundwater and surface waters (23). This aspect needs consideration in assessments of the risk associated with use of crops as green manure to replace chemical fertilizers and pesticides as well as in genetic engineering approaches to design crops with improved pest resistance.

DISCLOSURE STATEMENT The authors are not aware of any affiliations, memberships, funding, or financial holdings that might be perceived as affecting the objectivity of this review. 176

Gleadow

·

Møller

PP65CH06-Gleadow

ARI

8 April 2014

21:52

ACKNOWLEDGMENTS B.L.M. acknowledges financial support from the Villum Foundation to the VILLUM Research Center for Plant Plasticity, from the Novo Nordisk Foundation to the Plant Pathway Discovery Section at the Center for Biosustainability, and from the University of Copenhagen UCPH Excellence Programme for Interdisciplinary Research to Center for Synthetic Biology “bioSYNergy.” We gratefully acknowledge the support of the Australian Research Council (DP130101049 to B.L.M. and R.M.G.). We also thank Samantha Fromhold and Mohammed Saddik Motawia for help with figure preparation.


LITERATURE CITED 1. Adewusi SRA. 1990. Turnover of dhurrin in green sorghum seedlings. Plant Physiol. 94:1219–24 2. Agrawal AA. 2011. New synthesis—trade-offs in chemical ecology. J. Chem. Ecol. 37:230–31 3. Agrawal AA, Hastings AP, Johnson MTJ, Maron JL, Salminen J-P. 2012. Insect herbivores drive realtime ecological and evolutionary change in plant populations. Science 338:113–16 4. Ahn YO, Saino H, Mizutani M, Shimizu B, Sakata K. 2007. Vicianin hydrolase is a novel cyanogenic β-glycosidase specific to β-vicianoside (6-O-α-L-arabinopyranosyl-β-D-glucopyranoside) in seeds of Vicia angustifolia. Plant Cell Physiol. 48:938–39 5. Alves AAC, Setter TL. 2004. Abscisic acid accumulation and osmotic adjustment in cassava under water deficit. Environ. Exp. Bot. 51:259–71 6. Andersen MD, Busk PK, Svendsen I, Møller BL. 2000. Cytochromes P450 from cassava (Manihot esculenta Crantz) catalyzing the first steps in the biosynthesis of the cyanogenic glucosides linamarin and lotaustralin: cloning, functional expression in Pichia pastoris, and substrate specificity of the isolated recombinant enzymes. J. Biol. Chem. 275:1966–75 7. Bak S, Beisson F, Bishop G, Hamberger B, Hofer R, et al. 2011. Cytochromes P450. Arabidopsis Book ¨ 9:e0144 8. Bak S, Kahn RA, Nielsen HN, Møller BL, Halkier BA. 1998. Cloning of three A-type cytochromes P450, CYP71E1, CYP98, and CYP99 from Sorghum bicolor (L.) Moench by a PCR approach and identification by expression in Escherichia coli of CYP71E1 as a multifunctional cytochrome P450 in the biosynthesis of the cyanogenic glucoside dhurrin. Plant Mol. Biol. 36:393–405 9. Bak S, Paquette S, Morant M, Morant A, Saito S, et al. 2006. Cyanogenic glycosides: a case study for evolution and application of cytochromes P450. Phytochem. Rev. 5:309–29 10. Ballhorn DJ. 2011. Constraints of simultaneous resistance to a fungal pathogen and an insect herbivore in lima bean (Phaseolus lunatus L.). J. Chem. Ecol. 37:141–44 11. Ballhorn DJ, Heil M, Lieberei R. 2006. Phenotypic plasticity of cyanogenesis in lima bean Phaseolus lunatus—activity and activation of β-glucosidase. J. Chem. Ecol. 32:261–75 12. Ballhorn DJ, Kautz S, Jensen M, Schmitt I, Heil M, Hegeman AD. 2011. Genetic and environmental interactions determine plant defences against herbivores. J. Ecol. 99:313–26 13. Ballhorn DJ, Kautz S, Lion U, Heil M. 2008. Trade-offs between direct and indirect defences of lima bean (Phaseolus lunatus). J. Ecol. 96:971–80 14. Ballhorn DJ, Kautz S, Schädler M. 2013. Induced plant defense via volatile production is dependent on rhizobial symbiosis. Oecologia 172:833–46 15. Ballhorn DJ, Lieberei R, Ganzhorn JU. 2005. Plant cyanogenesis of Phaseolus lunatus and its relevance for herbivore-plant interaction: the importance of quantitative data. J. Chem. Ecol. 31:1445–73 16. Ballhorn DJ, Pietrowski A, Lieberei R. 2010. Direct trade-off between cyanogenesis and resistance to a fungal pathogen in lima bean (Phaseolus lunatus L.). J. Chem. Ecol. 98:226–36 17. Ballhorn DJ, Schiwy S, Jensen M, Heil M. 2008. Quantitative variability of direct chemical defense in primary and secondary leaves of lima bean (Phaseolus lunatus) and consequences for a natural herbivore. J. Chem. Ecol. 34:1298–301 18. Bari R, Jones JDG. 2009. Role of plant hormones in plant defence responses. Plant Mol. Biol. 69:473–88 19. Barnett RD, Caviness CE. 1968. Inheritance of hydrocyanic acid production in two sorghum × sudangrass crosses. Crop Sci. 8:89–91 www.annualreviews.org • Cyanogenic Glycosides

16. Shows that the trade-off between the production of CNglcs and volatile organic compounds results from the balance between relative rates of herbivory and pathogen attack.

177


PP65CH06-Gleadow

ARI

8 April 2014

21:52

20. Barros PM, Gonçalves N, Saibo NJM, Oliveira MM. 2012. Cold acclimation and floral development in almond bud break: insights into the regulatory pathways. J. Exp. Bot. 63:4585–96 21. Basile LJ, Willson RC, Sewell BT. 2008. Genome mining of cyanide-degrading nitrilases from filamentous fungi. Appl. Microbiol. Biotechnol. 80:427–35 22. Bazin A, Goverde M, Erhardt A, Shykoff JA. 2002. Influence of atmospheric carbon dioxide enrichment on induced response and growth compensation after herbivore damage in Lotus corniculatus. Ecol. Entomol. 27:271–78 23. Bjarnholt N, Laegdsmand M, Hansen HC, Jacobsen OH, Møller BL. 2008. Leaching of cyanogenic glucosides and cyanide from white clover green manure. Chemosphere 72:897–904 24. Bjarnholt N, Møller BL. 2008. Hydroxynitrile glucosides. Phytochemistry 69:1947–61 25. Blomstedt CK, Gleadow RM, O’Donnell N, Naur P, Jensen K, et al. 2012. A combined biochemical screen and TILLING approach identifies mutations in Sorghum bicolor L. Moench resulting in acyanogenic forage production. Plant Biotechnol. J. 10:54–66 26. Blumenthal SG, Hendrickson HR, Abrol YP, Conn EE. 1968. Cyanide metabolism in higher plants. III. The biosynthesis of β-cyanoalanine. J. Biol. Chem. 243:5302–7 27. Briggs MA. 1990. Chemical defense production in Lotus corniculatus L. I. The effects of nitrogen source on growth, reproduction and defense. Oecologia 83:27–31 28. Brown A. 2011. Effect of temperature and drought on the growth and nutritive value of cassava. Hon. Thesis, Monash Univ., Victoria, Aust. 29. Buchanan CD, Lim S, Salzman RA, Kagiampakis I, Morishige DT, et al. 2010. Sorghum bicolor’s transcriptome response to dehydration, high salinity and ABA. Plant Mol. Biol. 58:699–720 30. Burns AE, Gleadow RM, Woodrow IE. 2002. Light alters the allocation of nitrogen to cyanogenic glycosides in Eucalyptus cladocalyx. Oecologia 133:288–94 31. Burns AE, Gleadow RM, Zacarias A, Cuambe CE, Miller RE, Cavagnaro TR. 2012. Variations in the chemical composition of cassava (Manihot esculenta Crantz) leaves and roots as affected by genotypic and environmental variation. J. Agric. Food Chem. 60:4946–56 32. Busk PK, Møller BL. 2002. Dhurrin synthesis in sorghum is regulated at the transcriptional level and induced by nitrogen fertilization in older plants. Plant Physiol. 129:1222–31 33. Cavagnaro TR, Gleadow RM, Miller RE. 2011. Plant nutrient acquisition and utilisation in a high carbon dioxide world. Funct. Plant Biol. 38:87–96 34. Chassagne D, Crouzet JC, Bayonove CL, Baumes RL. 1996. Identification and quantification of passion fruit cyanogenic glycosides. J. Agric. Food Chem. 44:3817–20 35. Cicek M, Esen A. 1998. Structure and expression of a dhurrinase (β-glucosidase) from Sorghum. Plant Physiol. 116:1469–78 36. Clausen V, Frydenvang K, Koopmann R, Jørgensen LB, Abbiw DK, et al. 2002. Plant analysis by butterflies: occurrence of cyclopentenylglycines in Passifloraceae, Flacourtiaceae, and Turneraceae and discovery of the novel nonproteinogenic amino acid 2-(3 -cyclopentenyl)glycine in Rinorea. J. Nat. Prod. 65:542–47 37. Clegg DO, Conn EE, Janzen DH. 1979. Developmental fate of the cyanogenic glycoside linamarin in Costa Rican wild lima bean seeds. Nature 278:343–44 38. Coley PD, Bryant JP, Chapin FS III. 1985. Resource availability and plant antiherbivore defense. Science 230:895–99 39. Conn EE. 1980. Cyanogenic compounds. Annu. Rev. Plant Physiol. 31:433–51 40. Conn EE. 1991. The metabolism of a natural product: lessons learned from cyanogenic glycosides. Planta Med. 57:1–9 41. Conn EE, Seigler DS, Maslin BR, Dunn J. 1989. Cyanogenesis in Acacia subgenus Aculezferum. Phytochemistry 28:817–20 42. Cooper-Driver G, Finch S, Swain T. 1977. Seasonal variation in secondary plant compounds in relation to the palatability of Pteridium aqulinum. Biochem. Syst. Ecol. 5:177–83 43. Corkhill L. 1942. Cyanogenesis in white clover (Trifolium repens L.) V. The inheritance of cyanogenesis. N.Z. J. Sci. Technol. B 23:178–93 44. Cutler AJ, Conn EE. 1981. The biosynthesis of cyanogenic glucosides in Linum usitatissimum (linen flax) in vitro. Arch. Biochem. Biophys. 212:468–74

25. Uses ethyl methanesulfonate– generated sorghum mutants with altered cyanogenic status to provide a new crop model for study.

178

Gleadow

·

Møller


PP65CH06-Gleadow

ARI

8 April 2014

21:52

45. Dahler JM, McConchie C, Turnbull CGN. 1995. Quantification of cyanogenic glycosides in seedlings of three Macadamia (Proteaceae) species. Aust. J. Bot. 43:619–28 46. Dinh ST, Baldwin IT, Galis I. 2013. The HERBIVORE ELICITOR-REGULATED1 gene enhances abscisic acid levels and defenses against herbivores in Nicotiana attenuata plants. Plant Physiol. 162:2106– 24 47. Dugas DV, Monaco MK, Olsen A, Klein RR, Kumari S, et al. 2011. Functional annotation of the transcriptome of Sorghum bicolor in response to osmotic stress and abscisic acid. BMC Genomics 12:514 48. Elias M, Nambisan B, Sudhakaran PR. 1997. Characterization of linamarase of latex and its localization in petioles in cassava. Arch. Biochem. Biophys. 341:222–28 49. Erb M, Meldau S, Howe GA. 2012. Role of phytohormones in insect-specific plant reactions. Trends Plant Sci. 17:250–59 50. Ernesto M, Cardoso AP, Nicala D, Mirione E, Massaza F, et al. 2002. Persistent konzo and cyanogen toxicity from cassava in northern Mozambique. Acta Trop. 22:357–62 51. Finnie JW, Windsor PA, Kessell AE. 2011. Neurological diseases of ruminant livestock in Australia. II. Toxic disorders and nutritional deficiencies. Aust. Vet. J. 89:247–53 52. Flamatti GR, Waters MT, Scaffidi A, Merritt DJ, Ghisalberti EL, et al. 2013. Karrikin and cyanohydrin smoke signals provide clues to new endogenous plant signaling compounds. Mol. Plant 6:29–37 53. Forslund K, Morant M, Jørgensen B, Olsen CE, Asamizu E, et al. 2004. Biosynthesis of the nitrile glucosides rhodiocyanoside A and D and the cyanogenic glucosides lotaustralin and linamarin in Lotus japonicus. Plant Physiol. 135:71–84 54. Foulds W. 1982. Polymorphism for cyanogenesis in Lotus australis Andr. populations at Greenough Front Flats, Western Australia. Aust. J. Bot. 30:211–17 55. Foulds W, Grime JP. 1972. The response of cyanogenc and acyanogenic phenotypes of Trifolium repens to soil moisture supply. Heredity 28:181–87 56. Franks TK, Powell KS, Choimes S, Marsh E, Iocco P, et al. 2006. Consequences of transferring three sorghum genes for secondary metabolite (cyanogenic glucoside) biosynthesis to grapevine hairy roots. Transgenic Res. 15:181–95 57. Franks TK, Yadollahi A, Wirthensohn MG. 2008. A seed coat cyanohydrin glucosyltransferase is associated with bitterness in almond (Prunus dulcis) kernels. Funct. Plant Biol. 35:236–46 58. Frehner M, Conn EE. 1987. The linamarin β-glucosidase in Costa Rica wild bean (Phaseolus lunatus L.) is apoplastic. Plant Physiol. 84:1296–300 59. Frehner M, Luscher A, Hebeisen T, Zanetti S, Schubiger F, Scalet M. 1997. Effects of elevated partial pressure of carbon dioxide and season of the year on forage quality and cyanide concentration of Trifolium repens L. from a FACE experiment. Acta Oecologica 18:297–304 60. Gershenzon J. 1989. The cost of plant chemical defense against herbivory: a biochemical perspective. In Insect–Plant Interactions, ed. EA Bernays, pp. 105–73. Bacon Raton, FL: CRC 61. Gierl A, Frey M. 2001. Evolution of benzoxazinone biosynthesis and indole production in maize. Planta 213:493–98 62. Gleadow RM, Edwards E, Evans JR. 2009. Changes in nutritional value of cyanogenic Trifolium repens at elevated CO2 . J. Chem. Ecol. 35:476–47 63. Gleadow RM, Evans JR, McCaffery S, Cavagnaro TR. 2009. Growth and nutritive value of cassava (Manihot esculenta Cranz.) are reduced when grown in elevated CO2 . Plant Biol. 11:76–82 64. Gleadow RM, Foley WJ, Woodrow IE. 1998. Enhanced CO2 alters the relationship between photosynthesis and defence in cyanogenic Eucalyptus cladocalyx F. Muell. Plant Cell Environ. 21:12–22 65. Gleadow RM, Haburjak J, Dunn JE, Conn ME, Conn EE. 2008. Frequency and distribution of cyanogenic glycosides in Eucalyptus L’Hérit. Phytochemistry 69:1870–74 66. Gleadow RM, Møller BL, Cavagnaro T, Miller R, Stuart P, et al. 2013. Impacts of climate change on allocation of N to cyanogenic glycosides. Pharm. Biol. 50:623 (Abstr.) 67. Gleadow RM, Veechies AC, Woodrow IE. 2003. Cyanogenic Eucalyptus nobilis is polymorphic for both prunasin and specific β-glucosidases. Phytochemistry 63:699–704 68. Gleadow RM, Woodrow IE. 2000. Polymorphism in cyanogenic glycoside content and cyanogenic β-glucosidase activity in natural populations of Eucalyptus cladocalyx. Aust. J. Plant Physiol. 27:693–99 www.annualreviews.org • Cyanogenic Glycosides

179

PP65CH06-Gleadow

ARI

8 April 2014


70. Shows that the effectiveness of CNglcs as a general defense depends on HCNp, diet quality, and mode of feeding.

180

21:52

69. Gleadow RM, Woodrow IE. 2000. Temporal and spatial variation in cyanogenic glycosides in Eucalyptus cladocalyx. Tree Physiol. 20:591–98 70. Gleadow RM, Woodrow IE. 2002. Constraints on effectiveness of cyanogenic glycosides in herbivore defense. J. Chem. Ecol. 28:1301–13 71. Gleadow RM, Woodrow IE. 2002. Defense chemistry of cyanogenic Eucalyptus cladocalyx seedlings is affected by water supply. Tree Physiol. 22:939–45 72. Goodger JQD, Ades PK, Woodrow IE. 2004. Cyanogenesis in Eucalyptus polyanthemos seedlings: heritability, ontogeny and effect of soil nitrogen. Tree Physiol. 24:681–88 73. Goodger JQD, Capon RJ, Woodrow IE. 2002. Cyanogenic polymorphism in Eucalyptus polyanthemos Schauer subsp. vestita L. Johnson and K. Hill (Myrtaceae). Biochem. Syst. Ecol. 30:617–30 74. Goodger JQD, Gleadow RM, Woodrow IE. 2006. Growth cost and ontogenetic expression patterns of defence in cyanogenic Eucalyptus spp. Trees 20:757–65 75. Goodger JQD, Woodrow IE. 2002. Cyanogenic polymorphism as an indicator of genetic diversity in the rare species Eucalyptus yarraensis (Myrtaceae). Funct. Plant Biol. 29:1445–52 76. Gruhnert C, Biehl B, Selmar D. 1994. Compartmentation of cyanogenic glucosides and their degrading enzymes. Planta 195:36–42 77. Gwilt JR. 1960. Odour of potassium cyanide. Lancet 276:1197 78. Halkier BA, Møller BL. 1989. Biosynthesis of the cyanogenic glucoside dhurrin in seedlings of Sorghum bicolor (L.) Moench and partial purification of the enzyme system involved. Plant Physiol. 90:1552–59 79. Halkier BA, Møller BL. 1991. Involvement of cytochrome P-450 in the biosynthesis of dhurrin in Sorghum bicolor (L.) Moench. Plant Physiol. 96:10–17 80. Hansen KS, Kristensen C, Tattersall DB, Jones PR, Olsen CE, et al. 2003. The in vitro substrate regiospecificity of recombinant UGT85B1, the cyanohydrin glucosyltransferase from Sorghum bicolor. Phytochemistry 64:143–51 81. Hayden KJ, Parker IM. 2002. Plasticity in cyanogenesis of Trifolium repens L.: inducibility, fitness costs and variable expression. Evol. Ecol. Res. 4:155–68 82. Heraud P, Caine S, Sanson G, Gleadow RM, Wood BR, McNaughton D. 2007. Focal plane array infrared imaging: a new way to analyse leaf tissue. New Phytol. 173:216–25 83. Herms DA, Mattson WJ. 1992. The dilemma of plants: to grow or defend. Q. Rev. Biol. 67:283–335 84. Hickel A, Hasslacher M, Griengl H. 1996. Hydroxynitrile lyases: functions and properties. Physiol. Plant. 98:891–98 85. Hosel W, Berlin J, Hanzlik TN, Conn EE. 1987. In-vitro biosynthesis of 1-(4 -hydroxyphenol)-2¨ nitroethane and production of cyanogenic compounds in osmotically stressed cell suspension cultures of Eschscholtzia californica Cham. Planta 166:176–81 86. Hughes J, Decarvalho JPC, Hughes MA. 1994. Purification, characterization, and cloning of αhydroxynitrile lyase from cassava (Manihot esculenta Crantz). Arch. Biochem. Biophys. 311:496–502 87. Hughes MA. 1991. The cyanogenic polymorphism in Trifolium repens L. (white clover). Heredity 66:105– 15 88. Ibenthal WD, Pourmohseni H, Grosskopf S, Oldenburg H, Shafieiazad S. 1993. New approaches towards biochemical mechanisms of resistance susceptibility of Gramineae to powdery mildew (Erysiphe graminis). Angew. Bot. 67:97–106 89. Itkin M, Heinig U, Tzfadia O, Bhide AJ, Shinde B, et al. 2013. Biosynthesis of antinutritional alkaloids in solanaceous crops is mediated by clustered genes. Science 341:175–79 90. Jenrich R, Trompetter I, Bak SS, Olsen CE, Møller BL, Piotrowski M. 2007. Evolution of heteromic nitrilase complexes in Poaceae with new functions in nitrile metabolism. Proc. Natl. Acad. Sci. USA 104:18848–53 91. Jensen K, Osmani SA, Hamann T, Naur P, Møller B. 2011. Homology modeling of the three membrane proteins of the dhurrin metabolon: catalytic sites, membrane surface association and protein-protein interactions. Phytochemistry 72:2113–23 92. Jensen NB, Zagrobelny M, Hjernø K, Olsen CE, Houghton-Larsen J, et al. 2011. Convergent evolution in biosynthesis of cyanogenic defence compounds in plants and insects. Nat. Commun. 2:273 93. Jones DA. 1998. Why are so many food plants cyanogenic? Phytochemistry 47:155–62 Gleadow

·

Møller


PP65CH06-Gleadow

ARI

8 April 2014

21:52

94. Jones PR, Møller BL, Hoj PB. 1999. The UDP-glucose:p-hydroxymandelonitrile-O-glucosyltransferase that catalyzes the last step in synthesis of the cyanogenic glucoside dhurrin in Sorghum bicolor: isolation, cloning, heterologous expression, and substrate specificity. J. Biol. Chem. 274:35483–91 95. Jørgensen K, Bak S, Busk PK, Sørenson C, Olsen CE, et al. 2005. Cassava plants with a depleted cyanogenic glucoside content in leaves and tubers. Distribution of cyanogenic glucosides, their site of synthesis and transport, and blockage of the biosynthesis by RNA interference technology. Plant Physiol. 139:363–74 96. Jørgensen K, Morant AV, Morant M, Jensen NB, Olsen CE, et al. 2011. Biosynthesis of the cyanogenic glucosides linamarin and lotaustralin in cassava: isolation, biochemical characterization, and expression pattern of CYP71E7, the oxime-metabolizing cytochrome P450 enzyme. Plant Physiol. 155:282–92 97. Jørgensen K, Rasmussen AV, Morant M, Nielsen AH, Bjarnholt N, et al. 2005. Metabolon formation and metabolic channeling in the biosynthesis of plant natural products. Curr. Opin. Plant Biol. 8:280– 91 98. Kadow D, Voß K, Selmar D, Lieberei R. 2012. The cyanogenic syndrome in rubber tree Hevea brasiliensis: tissue-damage-dependent activation of linamarase and hydroxynitrile lyase accelerates hydrogen cyanide release. Ann. Bot. 109:1253–62 99. Kahn RA, Bak S, Svendsen I, Halkier BA, Møller BL. 1997. Isolation and reconstitution of cytochrome P450ox and in vitro reconstitution of the entire biosynthetic pathway of the cyanogenic glucoside dhurrin from sorghum. Plant Physiol. 115:1661–70 100. Kahn RA, Fahrendorf T, Halkier BA, Møller BL. 1999. Substrate specificity of the cytochrome P450 enzymes CYP79A1 and CYP71E1 involved in the biosynthesis of the cyanogenic glucoside dhurrin in Sorghum bicolor (L.) Moench. Arch. Biochem. Biophys. 363:9–18 101. Kakes P. 1985. Linamerase and other β-glucosidases are present in the cell walls of Trifolium repens L. leaves. Planta 166:156–60 102. Kakes P. 1989. An analysis of the costs and benefits of the cyanogenic system in Trifolium repens L. Theor. Appl. Genet. 77:111–18 103. Kakes P. 1994. The function of cyanogenesis in cassava. Acta Hortic. 375:79–85 104. Kakes P. 1997. Difference between the male and female components of fitness associated with the gene Ac in Trifolium repens. Acta Bot. Neerl. 46:219–23 105. Kannangara R, Motawia MS, Hansen NKK, Paquette SM, Olsen CE, et al. 2011. Characterization and expression profile of two UDP-glucosyltransferases, UGT85K4 and UGT85K5, catalyzing the last step in cyanogenic glucoside biosynthesis in cassava. Plant J. 68:287–301 106. Kempel A, Brandl R, Schädler M. 2009. Symbiotic soil microorganisms as players in aboveground plantherbivore interactions: the role of rhizobia. Oikos 118:634–40 107. Kerchev PL, Fenton B, Foyer CH, Hancock RD. 2012. Plant responses to insect herbivory: interactions between photosynthesis, reactive oxygen species and hormone signalling pathways. Plant Cell Environ. 35:441–53 108. King NLR, Bradbury JH. 1995. Bitterness of cassava: identification of a new apiosyl glucoside and other compounds that affect its bitter taste. J. Sci. Food Agric. 68:223–30 109. Koch BM, Sibbesen O, Halkier BA, Svendsen I, Møller BL. 1995. The primary sequence of cytochrome P450tyr, the multifunctional N-hydroxylase catalyzing the conversion of L-tyrosine to p-hydroxyphenylacetaldehyde oxime in the biosynthesis of the cyanogenic glucoside dhurrin in Sorghum bicolor (L.) Moench. Arch. Biochem. Biophys. 323:177–86 110. Koepke T, Schaeffer S, Harper A, Dicenta F, Edwards M, et al. 2013. Comparative genomics analysis in Prunoideae to identify biologically relevant polymorphisms. Plant Biotechnol. J. 11:883–93 111. Kojima M, Poulton JE, Thayer SS, Conn EE. 1979. Tissue distributions of dhurrin and of enzymes involved in its metabolism in leaves of Sorghum bicolor. Plant Physiol. 63:1022–28 112. Kongsawadworakul P, Viboonjun U, Romruensukharom P, Chantuma P, Ruderman S, Chrestin H. 2009. The leaf, inner bark and latex cyanide potential of Hevea brasiliensis: evidence for involvement of cyanogenic glucosides in rubber yield. Phytochemistry 70:730–39 113. Kristensen C, Morant M, Olsen CE, Ekstrom CT, Galbraith DW, et al. 2005. Metabolic engineering of dhurrin in transgenic Arabidopsis plants with marginal inadvertent effects on the metabolome and transcriptome. Proc. Natl. Acad. Sci. USA 102:1779–84 www.annualreviews.org • Cyanogenic Glycosides

98. Shows that some fungi increase HCNc by inducing catabolic enzyme activity to suppress natural phytoalexin responses.

181

ARI

8 April 2014

21:52

114. Lamont BB. 1993. Injury-induced cyanogenesis in vegetative and reproductive parts of two Grevillea species and their F1 hybrid. Ann. Bot. 71:537–42 115. Leavesley HB, Li L, Prabhakaran K, Borowitz JL, Ison GE. 2008. Interaction of cyanide and nitric oxide with cytochrome c oxidase: implications for acute cyanide toxicity. Toxicol. Sci. 101:101–11 116. Lee J, Zhang G, Wood E, Castillo CR, Mitchell AE. 2013. Quantification of amygdalin in nonbitter, semibitter, and bitter almonds (Prunus dulcis) by UHPLC-(ESI)QqQ MS/MS. J. Agric. Food Chem. 61:7754–59 117. Li B, Knudsen C, Hansen NK, Jørgensen K, Kannangara R, et al. 2013. Visualizing metabolite distribution and enzymatic conversion in plant tissues by desorption electrospray ionization mass spectrometry imaging. Plant J. 74:1059–71 118. Lieberei R, Selmar D, Biehl B. 1985. Metabolisation of cyanogenic glucosides in Hevea brasiliensis. Plant Syst. Evol. 150:49–63 119. Lieberei R. 2007. South American leaf blight of the rubber tree (Hevea spp.): new steps in plant domestication using physiological features and molecular markers. Ann. Bot. 100:1125–42 120. Loyd RC, Gray E. 1970. Amount and distribution of hydrocyanic acid potential during the life cycle of plants of three sorghum cultivars. Agron. J. 62:394–97 121. McBee GG, Miller FR. 1980. Hydrocyanid acid potential in several sorghum breeding lines as affected by nitrogen fertilization and variable harvests. Crop Sci. 20:232–35 122. McFarlane IJ, Lees EM, Conn EE. 1975. The in vitro biosynthesis of dhurrin, the cyanogenic glycoside of Sorghum bicolor. J. Biol. Chem. 250:4708–13 123. McKey D. 1974. Adaptive patterns in alkaloid physiology. Am. Nat. 108:305–20 124. McKey D, Cavagnaro TR, Cliff J, Gleadow RM. 2010. Chemical ecology in coupled human and natural systems: people, manioc, multitrophic interactions and global change. Chemoecology 20:109–33 125. McMahon JM, White WLB, Sayre RT. 1995. Cyanogenesis in cassava (Manihot esculenta Crantz). J. Exp. Bot. 46:731–41 126. Miller RE, Gleadow RM, Cavagnaro TR. 2014. Age versus stage: Does ontogeny modify the effect of phosphorus and arbuscular mycorrhizas on above- and below-ground defence in forage sorghum? Plant Cell Environ. 37:929–42 127. Miller RE, Gleadow RM, Woodrow IE. 2004. Cyanogenesis in tropical Prunus turneriana: characterisation, variation and response to low light. Funct. Plant Biol. 31:491–503 128. Miller RE, Jensen R, Woodrow IE. 2006. Frequency of cyanogenesis in tropical rainforests of Far North Queensland, Australia. Ann. Bot. 97:1017–44 129. Miller RE, McConville MJ, Woodrow IE. 2006. Cyanogenic glycosides from the rare Australian endemic rainforest tree Clerodendrum grayi (Lamiaceae). Phytochemistry 67:43–51 130. Miller RE, Simon J, Woodrow IE. 2006. Cyanogenesis in the Australian tropical rainforest endemic Brombya platynema (Rutaceae): chemical characterisation and polymorphism. Funct. Plant Biol. 33:477– 86 131. Miller RE, Tuck KL. 2013. Reports on the distribution of aromatic cyanogenic glycosides in Australian tropical rainforest tree species of the Lauraceae and Sapindaceae. Phytochemistry 92:146–52 132. Møller BL. 2010. Dynamic metabolons. Science 330:1328–29 133. Møller BL. 2010. Functional diversifications of cyanogenic glucosides. Curr. Opin. Plant Biol. 13:337–46 134. Møller BL, Conn EE. 1979. The biosynthesis of cyanogenic glucosides in higher plants: Nhydroxytyrosine as an intermediate in the biosynthesis of dhurrin by Sorghum bicolor (L.) Moench. J. Biol. Chem. 254:8575–83 135. Moore BP. 1967. Hydrogen cyanide in the defensive secretions of larval Paropsini (Coleoptera: Chrysomelidae). Aust. J. Entomol. 6:36–38 136. Morant AV, Bjarnholt N, Kragh ME, Kjaergaard CH, Jorgensen K, et al. 2008. The β-glucosidases responsible for bioactivation of hydroxynitrile glucosides in Lotus japonicus. Plant Physiol. 147:1072–91 137. Morant AV, Jørgensen K, Jørgensen C, Paquette SM, Sánchez-Pérez R, et al. 2008. β-glucosidases as detonators of plant chemical defense. Phytochemistry 69:1795–813 138. Mugford ST, Louveau T, Melton R, Qi X, Bakht S, et al. 2013. Modularity of plant metabolic gene clusters: a trio of linked genes that are collectively required for acylation of triterpenes in oat. Plant Cell 25:1078–92


PP65CH06-Gleadow

182

Gleadow

·

Møller


PP65CH06-Gleadow

ARI

8 April 2014

21:52

139. Neilson EH, Goodger JQD, Motawia MS, Bjarnholt N, Frisch T, et al. 2011. Phenylalanine derived cyanogenic diglucosides from Eucalyptus camphora and their abundances in relation to ontogeny and tissue type. Phytochemistry 72:2325–34 140. Neilson EH, Goodger JQD, Woodrow IE, Møller BL. 2013. Plant chemical defence: at what cost? Trends Plant Sci. 18:250–58 141. Nielsen AZ, Ziersen B, Jensen K, Lassen LM, Olsen CE, et al. 2013. Redirecting photosynthetic reducing power toward bioactive natural product synthesis. ACS Synth. Biol. 2:308–15 142. Nielsen JS, Møller BL. 1999. Biosynthesis of cyanogenic glucosides in Triglochin maritima and the involvement of cytochrome P450 enzymes. Arch. Biochem. Biophys. 368:121–30 143. Nielsen KA, Olsen CE, Pontoppidan K, Møller BL. 2002. Leucine-derived cyanoglucosides in barley. Plant Physiol. 129:1066–75 144. Nielsen KA, Tattersall DB, Jones PR, Møller BL. 2008. Metabolon formation in dhurrin biosynthesis. Phytochemistry 69:88–98 145. Nishida R. 2002. Sequestration of defensive substances from plants by Lepidoptera. Annu. Rev. Entomol. 47:57–92 146. O’Donnell NH. 2012. Regulation of cyanogenesis in forage sorghum. PhD Thesis, Monash Univ., Victoria, Aust. 147. O’Donnell NH, Møller BL, Neale AD, Hamill JD, Blomstedt CK, Gleadow RM. 2013. Effects of PEG-induced osmotic stress on growth and dhurrin levels of forage sorghum. Plant Physiol. Biochem. 73:83–92 148. Oke OL. 1969. The role of hydrocyanic acid in nutrition. World Rev. Nutr. Diet. 11:170–98 149. Olafsdottir ES, Jorgensen LB, Jaroszewski JW. 1992. Substrate specificity in the biosynthesis of cyclopentanoid cyanohydrin glucosides. Phytochemistry 31:4129–34 150. Olsen KM, Hsu S-C, Small LL. 2008. Evidence on the molecular basis of the Ac/ac adaptive cyanogenesis polymorphism in white clover (Trifolium repens L.). Genetics 179:517–26 151. Olsen KM, Kooyers NJ, Small LL. 2013. Recurrent gene deletions and the evolution of adaptive cyanogenesis polymorphisms in white clover (Trifolium repens L.). Mol. Ecol. 22:724–38 152. Olsen KM, Sutherland BL, Small LL. 2007. Molecular evolution of the Li/li chemical defence polymorphism in white clover (Trifolium repens L.). Mol. Ecol. 16:4180–93 153. Osbourn AE. 1996. Preformed antimicrobial compounds and plant defense against fungal attack. Plant Cell 8:1821–31 154. Patel CJ, Wright MJ. 1958. The effect of certain nutrients upon the hydrocyanic acid content of sudangrass grown in nutrient solution. Agron. J. 50:645–47 155. Peterson SC. 1986. Breakdown products of cyanogenesis repellency and toxicity to predatory ants. Naturwissenschaften 73:627–28 156. Peterson SC, Johnson ND, LeGuyader JL. 1987. Defensive regurgitation of allelochemicals derived from host cyanogenesis by eastern tent caterpillars. Ecology 68:1268–72 157. Piotrowski M. 2008. Primary or secondary? Versatile nitrilases in plant metabolism. Phytochemistry 69:2655–67 158. Piotrowski M, Volmer J. 2006. Cyanide metabolism in higher plants: cyanoalanine hydratase is a NIT4 homolog. Plant Mol. Biol. 61:111–22 159. Poulton JE, Li CP. 1994. Tissue level compartmentation of (R)-amygdalin and amygdalin hydrolase prevents large-scale cyanogenesis in undamaged Prunus seeds. Plant Physiol. 104:29–35 160. Pourmohseni H, Ibenthal WD. 1991. Novel β-cyanoglucosides in the epidermal tissue of barley and their possible role in the barley-powdery mildew interaction. Angew. Bot. 65:341–50 161. Qi X, Bakht S, Leggett M, Maxwell C, Melton R, Osbourn A. 2004. A gene cluster for secondary metabolism in oat: implications for the evolution of metabolic diversity in plants. Proc. Natl. Acad. Sci. USA 101:8233–38 162. Richmond GS, Ghisalberti EL. 1994. Seed dormancy and germination mechanisms in Eremophila (Myoporaceae). Aust. J. Bot. 42:705–15 163. Rockenbach J, Nahrstedt A, Wray V. 1992. Cyanogenic glycosides from Psydrax and Oxyanthus species. Phytochemistry 31:567–70 www.annualreviews.org • Cyanogenic Glycosides

140. Shows that plants reduce defense costs through the use of shared biosynthetic pathways and integration into the primary metabolism.

151. Provides a molecular analysis of Ac and Li alleles in clover that shows recent evolution but no evidence for balancing selection.

153. Reviews the function of CNglcs as a defense against pathogens.

183


PP65CH06-Gleadow

ARI

8 April 2014

178. Proposes that enhanced synthesis of plant secondary products under stressful conditions provides a mechanism for dissipating excess excitation energy.

187. Shows that genes encoding CNglc synthesis in different species are clustered, promoting stable inheritance of functional pathways.

184

21:52

164. Salzman RA, Brady JA, Finlayson SA, Buchanan CD, Summer EJ, et al. 2005. Transcriptional profiling of sorghum induced by methyl jasmonate, salicylic acid, and aminocyclopropane carboxylic acid reveals cooperative regulation and novel gene responses. Plant Physiol. 138:352–68 165. Sánchez-Pérez R, Belmonte FS, Borch J, Dicenta F, Møller BL, Jørgensen K. 2012. Prunasin hydrolases during fruit development in sweet and bitter almonds. Plant Physiol. 158:1916–32 166. Sánchez-Pérez R, Jørgensen K, Motawia MS, Dicenta F, Møller BL. 2009. Tissue and cellular localization of individual β-glycosidases using a substrate-specific sugar reducing assay. Plant J. 60:894–906 167. Sánchez-Pérez R, Jørgensen K, Olsen CE, Dicenta F, Møller BL. 2008. Bitterness in almonds. Plant Physiol. 146:1040–52 168. Santner A, Calderon-Villalobos LI, Estelle M. 2009. Plant hormones are versatile chemical regulators of plant growth. Nat. Chem. Biol. 5:301–7 169. Saunders JA, Conn EE. 1978. Presence of the cyanogenic glucoside dhurrin in isolated vacuoles from sorghum. Plant Physiol. 61:154–57 170. Schappert PJ, Shore JS. 1994. Cyanogenesis in Turnera ulmifolia L. (Turneraceae). I. Phenotypic distribution and genetic variation for cyanogenesis on Jamaica. Heredity 74:392–404 171. Schappert PJ, Shore JS. 1999. Cyanogenesis, herbivory and plant defense in Turnera ulmifolia on Jamaica. Ecoscience 6:511–20 172. Schappert PJ, Shore JS. 1999. Effects of cyanogenesis polymorphism in Turnera ulmifolia on Euptoieta hegesia and potential anolis predators. J. Chem. Ecol. 25:1455–79 173. Schappert PJ, Shore JS. 2000. Cyanogenesis in Turnera ulmifolia L. (Turneraceae): II. Developmental expression, heritability and cost of cyanogenesis. Evol. Ecol. Res. 2:337–52 174. Schwarz B, Wray V, Proksch P. 1996. A cyanogenic glycoside from Canthium schimperianum. Phytochemistry 42:633–36 175. Schwind P, Wray V, Nahrstedt A. 1990. Structure elucidation of an acylated cyanogenic triglycoside, and further cyanogenic constituents from Xeranthemum cylindraceum. Phytochemistry 29:1903–11 176. Selmar D. 1993. Transport of cyanogenic glucosides: linustatin uptake by Hevea cotyledons. Planta 191:191–99 177. Selmar D. 2010. Biosynthesis of cyanogenic glycosides, glucosinolates and non-protein amino acids. Annu. Plant Rev. 40:92–181 178. Selmar D, Kleinwachter M. 2013. Stress enhances the synthesis of secondary plant products: ¨ the impact of stress-related over-reduction on the accumulation of natural products. Plant Cell Physiol. 54:817–26 179. Selmar D, Lieberei R, Biehl B. 1988. Mobilization and utilization of cyanogenic glycosides: the linustatin pathway. Plant Physiol. 86:711–16 180. Sendker J, Nahrstedt A. 2009. Generation of primary amide glucosides from cyanogenic glucosides. Phytochemistry 70:388–93 181. Seo S, Mitsuhara I, Feng J, Iwai J, Hasegawa M, Ohashi Y. 2011. Cyanide, a coproduct of plant hormone ethylene biosynthesis, contributes to the resistance of rice to blast fungus. Plant Physiol. 155:502–14 182. Shimada M, Conn EE. 1977. The enzymatic conversion of p-hydroxyphenylacetaldoxime to p-hydroxymandelonitrile. Arch. Biochem. Biophys. 180:199–207 183. Sibbesen O, Koch B, Halkier BA, Møller BL. 1995. Cytochrome P-450TYR is a multifunctional hemethiolate enzyme catalyzing the conversion of L-tyrosine to p-hydroxyphenylacealdehyde oxime in the biosynthesis of the cyanogenic glucoside dhurrin in Sorghum bicolor (L.) Moench. J. Biol. Chem. 270:3506– 11 184. Simon J, Gleadow RM, Woodrow IE. 2010. Allocation of nitrogen to chemical defence and plant functional traits is constrained by soil N. Tree Physiol. 30:1111–17 185. Stochmal A, Oleszek W. 1997. Changes of cyanogenic glucosides in white clover (Trifolium repens L.) during the growing season. J. Agric. Food Chem. 45:4333–36 186. Swain E, Li CP, Poulton JE. 1992. Tissue and subcellular localization of enzymes catabolizing (R)-amygdalin in mature Prunus serotina seeds. Plant Physiol. 100:291–300 187. Takos AM, Knudsen C, Lai D, Kannangara R, Motawia MS, et al. 2011. Genomic clustering of cyanogenic glucoside biosynthetic genes aids their identification in Lotus japonicus and suggests the repeated evolution of this chemical defence pathway. Plant J. 68:273–86 Gleadow

·

Møller


PP65CH06-Gleadow

ARI

8 April 2014

21:52

188. Takos AM, Lai D, Mikkelsen L, Maher AH, Shelton D, et al. 2010. Genetic screening identifies cyanogenesis-deficient mutants of Lotus japonicus and reveals enzymatic specificity in hydroxynitrile glucoside metabolism. Plant Cell 22:1605–19 189. Takos AM, Rook F. 2012. Why biosynthetic genes for chemical defense compounds cluster. Trends Plant Sci. 17:383–88 190. Tattersall DB, Bak S, Jones PR, Olsen CE, Nielsen JK, et al. 2001. Resistance to an herbivore through engineered cyanogenic glucoside synthesis. Science 293:1826–28 191. Thayer SS, Conn EE. 1981. Subcellular localisation of dhurrin β-glucosidase and hydroxynitrile lyase in the mesophyll cells of sorghum leaf blades. Plant Physiol. 67:617–22 192. Thygesen LG, Jørgensen K, Møller BL, Engelsen SB. 2004. Raman spectroscopic analysis of cyanogenic glucosides in plants: development of a flow injection surface-enhanced Raman scatter (FI-SERS) method for determination of cyanide. Appl. Spectrosc. 58:212–17 193. Till I. 1987. Variability of expression of cyanogenesis in white clover (Trifolium repens L.). Heredity 59:265–71 194. Vandegeer R, Miller RE, Bain M, Gleadow RM, Cavagnaro TR. 2013. Drought adversely affects tuber development and nutritional quality of the staple crop cassava (Manihot esculenta Crantz). Funct. Plant Biol. 40:195–200 195. Vickery PJ, Wheeler JL, Mulcahy C. 1987. Factors affecting the hydrogen cyanide potential of white clover (Trifolium repens L.). Aust. J. Agric. Res. 38:1053–59 196. Wang Q, Hillwig ML, Peters RJ. 2011. CYP99A3: functional identification of a diterpene oxidase from the momilactone biosynthetic gene cluster in rice. Plant J. 65:87–95 197. Webber BL, Rentz DCF, Ueshima N, Woodrow IE. 2003. Leucopodoptera eumundii, a new genus and species of katydid from the tropical rainforests of North Queensland, Australia (Orthoptera: Tettigoniidae: Phaneropterinae: Holochlorini). J. Orthoptera Res. 12:79–88 198. Webber BL, Woodrow IE. 2008. Intra-plant variation in cyanogenesis and the continuum of foliar plant defense traits in the rainforest tree Ryparosa kurrangii (Achariaceae). Tree Physiol. 28:977–84 199. Webber BL, Woodrow IE. 2009. Chemical and physical plant defence across multiple ontogenetic stages in a tropical rain forest understorey tree. J. Ecol. 97:761–71 200. Wheeler JL, Mulcahy C, Walcott JJ, Rapp GG. 1990. Factors affecting the hydrogen cyanide potential of forage sorghum. Aust. J. Agric. Res. 41:1093–100 201. Winzer T, Gazda V, He Z, Kaminski F, Kern M, et al. 2012. A Papaver somniferum 10-gene cluster for synthesis of the anticancer alkaloid noscapine. Science 336:1704–8 202. Zagrobelny M, Bak S, Møller BL. 2008. Cyanogenesis in plants and arthropods. Phytochemistry 69:1457– 68 203. Zagrobelny M, Bak S, Olsen CE, Møller BL. 2007. Intimate roles for cyanogenic glucosides in the life cycle of Zygaena filipendulae (Lepidoptera, Zygaenidae). Insect Biochem. Mol. Biol. 37:1189–97 204. Zagrobelny M, Bak S, Rasmussen AV, Jorgensen B, Naumann CM, Møller BL. 2004. Cyanogenic glucosides and plant–insect interactions. Phytochemistry 65:293–306 205. Zagrobelny M, Møller BL. 2011. Cyanogenic glucosides in the biological warfare between plants and insects: the Burnet moth-birdsfoot trefoil model system. Phytochemistry 72:1585–92 206. Zagrobelny M, Motawia MS, Olsen CE, Bak S, Møller BL. 2013. Male-to-female transfer of 5-hydroxytryptophan glucoside during mating in Zygaena filipendulae (Lepidoptera). Insect Biochem. Mol. Biol. 43:1037–44 agg J, Jørgensen K, et al. 2014. Sequestration, tis207. Zagrobelny M, Olsen CE, Pentzold S, Furstenberg-H¨ ¨ sue distribution and developmental transmission of cyanogenic glucosides in a specialist insect herbivore. Insect Biochem. Mol. Biol. 44:44–53 208. Zhu-Salzman K, Salzman RA, Ahn J-E, Koiwa H. 2004. Transcriptional regulation of sorghum defense determinants against a phloem-feeding aphid. Plant Physiol. 134:420–31

www.annualreviews.org • Cyanogenic Glycosides

185

PP65-FrontMatter

ARI

8 April 2014

Annual Review of Plant Biology Volume 65, 2014

23:6

Contents


Our Eclectic Adventures in the Slower Eras of Photosynthesis: From New England Down Under to Biosphere 2 and Beyond Barry Osmond p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 1 Sucrose Metabolism: Gateway to Diverse Carbon Use and Sugar Signaling Yong-Ling Ruan p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p33 The Cell Biology of Cellulose Synthesis Heather E. McFarlane, Anett Döring, and Staffan Persson p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p69 Phosphate Nutrition: Improving Low-Phosphate Tolerance in Crops Damar Lizbeth López-Arredondo, Marco Antonio Leyva-González, Sandra Isabel González-Morales, José López-Bucio, and Luis Herrera-Estrella p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p95 Iron Cofactor Assembly in Plants Janneke Balk and Theresia A. Schaedler p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 125 Cyanogenic Glycosides: Synthesis, Physiology, and Phenotypic Plasticity Roslyn M. Gleadow and Birger Lindberg Møller p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 155 Engineering Complex Metabolic Pathways in Plants Gemma Farré, Dieter Blancquaert, Teresa Capell, Dominique Van Der Straeten, Paul Christou, and Changfu Zhu p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 187 Triterpene Biosynthesis in Plants Ramesha Thimmappa, Katrin Geisler, Thomas Louveau, Paul O’Maille, and Anne Osbourn p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 225 To Gibberellins and Beyond! Surveying the Evolution of (Di)Terpenoid Metabolism Jiachen Zi, Sibongile Mafu, and Reuben J. Peters p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 259 Regulation and Dynamics of the Light-Harvesting System Jean-David Rochaix p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 287 Gene Expression Regulation in Photomorphogenesis from the Perspective of the Central Dogma Shu-Hsing Wu p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 311 viii

PP65-FrontMatter

ARI

8 April 2014

23:6

Light Regulation of Plant Defense Carlos L. Ballaré p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 335 Heterotrimeric G Protein–Coupled Signaling in Plants Daisuke Urano and Alan M. Jones p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 365 Posttranslationally Modified Small-Peptide Signals in Plants Yoshikatsu Matsubayashi p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 385 Pentatricopeptide Repeat Proteins in Plants Alice Barkan and Ian Small p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 415


Division and Dynamic Morphology of Plastids Katherine W. Osteryoung and Kevin A. Pyke p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 443 The Diversity, Biogenesis, and Activities of Endogenous Silencing Small RNAs in Arabidopsis Nicolas G. Bologna and Olivier Voinnet p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 473 The Contributions of Transposable Elements to the Structure, Function, and Evolution of Plant Genomes Jeffrey L. Bennetzen and Hao Wang p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 505 Natural Variations and Genome-Wide Association Studies in Crop Plants Xuehui Huang and Bin Han p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 531 Molecular Control of Grass Inflorescence Development Dabing Zhang and Zheng Yuan p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 553 Male Sterility and Fertility Restoration in Crops Letian Chen and Yao-Guang Liu p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 579 Molecular Control of Cell Specification and Cell Differentiation During Procambial Development Kaori Miyashima Furuta, Eva Hellmann, and Ykä Helariutta p p p p p p p p p p p p p p p p p p p p p p p p p p 607 Adventitious Roots and Lateral Roots: Similarities and Differences Catherine Bellini, Daniel I. Pacurar, and Irene Perrone p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 639 Nonstructural Carbon in Woody Plants Michael C. Dietze, Anna Sala, Mariah S. Carbone, Claudia I. Czimczik, Joshua A. Mantooth, Andrew D. Richardson, and Rodrigo Vargas p p p p p p p p p p p p p p p p p p p 667 Plant Interactions with Multiple Insect Herbivores: From Community to Genes Jeltje M. Stam, Anneke Kroes, Yehua Li, Rieta Gols, Joop J.A. van Loon, Erik H. Poelman, and Marcel Dicke p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 689 Genetic Engineering and Breeding of Drought-Resistant Crops Honghong Hu and Lizhong Xiong p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 715

Contents

ix

PP65-FrontMatter

ARI

8 April 2014

23:6

Plant Molecular Pharming for the Treatment of Chronic and Infectious Diseases Eva Stoger, Rainer Fischer, Maurice Moloney, and Julian K.-C. Ma p p p p p p p p p p p p p p p p p p p 743 Genetically Engineered Crops: From Idea to Product Jose Rafael Prado, Gerrit Segers, Toni Voelker, Dave Carson, Raymond Dobert, Jonathan Phillips, Kevin Cook, Camilo Cornejo, Josh Monken, Laura Grapes, Tracey Reynolds, and Susan Martino-Catt p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 769 Errata


An online log of corrections to Annual Review of Plant Biology articles may be found at http://www.annualreviews.org/errata/arplant

x

Contents

Annual Reviews It’s about time. Your time. It’s time well spent.

New From Annual Reviews:

Annual Review of Statistics and Its Application Volume 1 • Online January 2014 • http://statistics.annualreviews.org

Editor: Stephen E. Fienberg, Carnegie Mellon University


Associate Editors: Nancy Reid, University of Toronto Stephen M. Stigler, University of Chicago The Annual Review of Statistics and Its Application aims to inform statisticians and quantitative methodologists, as well as all scientists and users of statistics about major methodological advances and the computational tools that allow for their implementation. It will include developments in the field of statistics, including theoretical statistical underpinnings of new methodology, as well as developments in specific application domains such as biostatistics and bioinformatics, economics, machine learning, psychology, sociology, and aspects of the physical sciences.

Complimentary online access to the first volume will be available until January 2015. table of contents:

• What Is Statistics? Stephen E. Fienberg • A Systematic Statistical Approach to Evaluating Evidence from Observational Studies, David Madigan, Paul E. Stang, Jesse A. Berlin, Martijn Schuemie, J. Marc Overhage, Marc A. Suchard, Bill Dumouchel, Abraham G. Hartzema, Patrick B. Ryan

• High-Dimensional Statistics with a View Toward Applications in Biology, Peter Bühlmann, Markus Kalisch, Lukas Meier • Next-Generation Statistical Genetics: Modeling, Penalization, and Optimization in High-Dimensional Data, Kenneth Lange, Jeanette C. Papp, Janet S. Sinsheimer, Eric M. Sobel

• The Role of Statistics in the Discovery of a Higgs Boson, David A. van Dyk

• Breaking Bad: Two Decades of Life-Course Data Analysis in Criminology, Developmental Psychology, and Beyond, Elena A. Erosheva, Ross L. Matsueda, Donatello Telesca

• Brain Imaging Analysis, F. DuBois Bowman

• Event History Analysis, Niels Keiding

• Statistics and Climate, Peter Guttorp

• Statistical Evaluation of Forensic DNA Profile Evidence, Christopher D. Steele, David J. Balding

• Climate Simulators and Climate Projections, Jonathan Rougier, Michael Goldstein • Probabilistic Forecasting, Tilmann Gneiting, Matthias Katzfuss • Bayesian Computational Tools, Christian P. Robert • Bayesian Computation Via Markov Chain Monte Carlo, Radu V. Craiu, Jeffrey S. Rosenthal • Build, Compute, Critique, Repeat: Data Analysis with Latent Variable Models, David M. Blei • Structured Regularizers for High-Dimensional Problems: Statistical and Computational Issues, Martin J. Wainwright

• Using League Table Rankings in Public Policy Formation: Statistical Issues, Harvey Goldstein • Statistical Ecology, Ruth King • Estimating the Number of Species in Microbial Diversity Studies, John Bunge, Amy Willis, Fiona Walsh • Dynamic Treatment Regimes, Bibhas Chakraborty, Susan A. Murphy • Statistics and Related Topics in Single-Molecule Biophysics, Hong Qian, S.C. Kou • Statistics and Quantitative Risk Management for Banking and Insurance, Paul Embrechts, Marius Hofert

Access this and all other Annual Reviews journals via your institution at www.annualreviews.org.

Annual Reviews | Connect With Our Experts Tel: 800.523.8635 (us/can) | Tel: 650.493.4400 | Fax: 650.424.0910 | Email: [email protected]

Mosaic physiology from developmental noise: within-organism physiological diversity as an alternative to phenotypic plasticity and phenotypic flexibility.

The ubiquity of phenotypic plasticity in plants: a synthesis.

Juxtaglomerular Cell Phenotypic Plasticity.

Straightforward rapid spectrophotometric quantification of total cyanogenic glycosides in fresh and processed cassava products.

[Correlation of cyanogenic glycosides with content of benzyl alcohol in cherry brandies (author's transl)].

Synthesis of episilon-rhodomycinone glycosides.

Plasticity of coral physiology under ocean acidification.

The higher the better? Differences in phenolics and cyanogenic glycosides in Sambucus nigra leaves, flowers and berries from different altitudes.

The Genetics of Phenotypic Plasticity. XIV. Coevolution.

Regulatory mechanisms link phenotypic plasticity to evolvability.

The phenotypic plasticity of developmental modules.

Endocrine regulation of predator-induced phenotypic plasticity.

Physiology and evolution at the crossroads of plasticity and inheritance.

Beyond risk, resilience, and dysregulation: phenotypic plasticity and human development.

Constraints on the evolution of phenotypic plasticity: limits and costs of phenotype and plasticity.

Enzymatic synthesis of epothilone A glycosides.

Functional and Phenotypic Plasticity of CD4(+) T Cell Subsets.

Phenotypic Plasticity and Selection: Nonexclusive Mechanisms of Adaptation.

Evolution of evolvability and phenotypic plasticity in virtual cells.

Genetic Regulation of Phenotypic Plasticity and Canalisation in Yeast Growth.

Lineage stability and phenotypic plasticity of Foxp3⁺ regulatory T cells.

The genetics of phenotypic plasticity. XII. Temporal and spatial heterogeneity.

Phase-transfer-catalyzed synthesis of flavonoid glycosides.

Enzymic synthesis of HexNAc-containing disaccharide glycosides.